TURUN YLIOPISTON JULKAISUJA
ANNALES UNIVERSITATIS TURKUENSIS
SARJA - SER. A I OSA - TOM. 422
ASTRONOMICA - CHEMICA - PHYSICA - MATHEMATICA
NADPH-DePeNDeNt tHioreDoxiN
SyStem iN regulAtioN of
CHloroPlASt fuNCtioNS
by
Anna Lepistö
TURUN YLIOPISTO
UNIVERSITY OF TURKU
Turku 2011
From the Laboratory of Molecular Plant Biology
Department of Biochemistry and Food Chemistry
University of Turku
FI-20014 Turku, Finland
Supervised by
Professor Eevi Rintamäki
Laboratory of Molecular Plant Biology
Department of Biochemistry and Food Chemistry
University of Turku
FI-20014 Turku, Finland
Reviewed by
Professor Elina Oksanen
Department of Biology
University of Eastern Finland
FI-80101 Joensuu, Finland
and
Professor Anja Hohtola
Department of Biology
University of Oulu
FI-90014 Oulu, Finland
Opponent
Professor Francisco Javier Cejudo
Department of Biology
University of Seville
ES-41092 Seville, Spain
ISBN 978-951-29-4694-5 (PRINT)
ISBN 978-951-29-4695-2 (PDF)
ISSN 0082-7002
Painosalama Oy – Turku, Finland 2011
LIST OF ORIGINAL PUBLICATIONS
This thesis is based on the following publications, which are referred to by their
Roman numerals in the text.
I
Lepistö A*, Kangasjärvi S*, Luomala EM, Brader G, Sipari N, Keränen M,
Keinänen M, Rintamäki E. 2009: Chloroplast NADPH thioredoxin reductase
interacts with photoperiodic development in Arabidopsis thaliana. Plant
Physiology 149:1261-1276
II
Lepistö A, Toivola J, Vignols F, Rintamäki E. 2011: Regulation of chloroplast
biogenesis by chloroplast NADPH-dependent thioredoxin system. Manuscript.
III
Lepistö A, Pakula E, Toivola J, Krieger-Liszkay A, Vignols F, Rintamäki E.
2011: Redox-regulation of starch and antioxidant metabolism by chloroplast
NADPH thioredoxin reductase in Arabidopsis grown under various
photoperiods. Manuscript.
IV
Kangasjärvi S, Lepistö A, Hännikäinen K, Piippo M, Luomala EM, Aro EM,
Rintamäki E. 2008: Diverse roles for chloroplast stromal and thylakoid-bound
ascorbate peroxidases in plant stress responses. Biochem J 412:275-285
*Equal contributions
Publication I has been reprinted by kind permission of American Society of Plant
Biologists. Copyright American Society of Plant Biologists.
Publication IV has been reprinted by kind permission of the Biochemical Society.
ABBREVIATIONS
AGPase
ALA
APX
ATP
CAO
CAT
CHLH, CHLI, CHLD
CRY
Cyt
DAHP
DHA
DSP4
FAD
FNR
FTR
G3P
GPX
GSH
GSSG
GUN4
HEMA1
LHC
MDA
NADPH
NTR
NTRC
P680, P700
Pi
PC
PCR
PGA
PHY
PLB
POR
PQ
Prx
PS
ROS
SOD
T-DNA
tRNA
ADP-glucose pyrophosphorylase
aminolevulinic acid
ascorbate peroxidase
adenosine triphosphate
chlorophyllide a oxygenase
catalase
subunits of Mg-chelatase
cryptochrome
cytochrome
3-deoxy-d-arabino-heptulosonate-7-phosphate
dehydroascorbate
dual specificity protein phosphatase
flavin adenine dinucleotide
ferredoxin-NADP+ oxidoreductase
ferredoxin-thioredoxin reductase
glyceraldehyde-3-phosphate
glutathione peroxidase
glutathione (reduced)
glutathione (oxidized)
regulator of Mg-chelatase
glutamyl-tRNA reductase
light-harvesting complex
monodehydroascorbate
nicotinamine adenine dinucleotide phosphate
NADPH-dependent thioredoxin reductase
plastid-localized NADPH-dependent thioredoxin
reductase C
reaction center chlorophyll of PSII and PSI, respectively
inorganic phosphate
plastocyanin
polymerase chain reaction
3-phosphoglycerate
phytocrome
prolamellar body
NADPH:protochlorophyllide oxidoreductase
plastoquinone
peroxiredoxin
photosystem
reactive oxygen species
superoxide dismutase
transfer DNA
transfer RNA
Trx
thioredoxin
ABSTRACT
Photosynthesis, the process in which carbon dioxide is converted into sugars using the
energy of sunlight, is vital for heterotrophic life on Earth. In plants, photosynthesis
takes place in specific organelles called chloroplasts. During chloroplast biogenesis,
light is a prerequisite for the development of functional photosynthetic structures. In
addition to photosynthesis, a number of other metabolic processes such as nitrogen
assimilation, the biosynthesis of fatty acids, amino acids, vitamins, and hormones are
localized to plant chloroplasts. The biosynthetic pathways in chloroplasts are tightly
regulated, and especially the reduction/oxidation (redox) signals play important roles in
controlling many developmental and metabolic processes in chloroplasts. Thioredoxins
are universal regulatory proteins that mediate redox signals in chloroplasts. They are
able to modify the structure and function of their target proteins by reduction of
disulfide bonds. Oxidized thioredoxins are restored via the action of thioredoxin
reductases. Two thioredoxin reductase systems exist in plant chloroplasts, the NADPHdependent thioredoxin reductase C (NTRC) and ferredoxin-thioredoxin reductase
(FTR). The ferredoxin-thioredoxin system that is linked to photosynthetic light
reactions is involved in light-activation of chloroplast proteins. NADPH can be
produced via both the photosynthetic electron transfer reactions in light, and in
darkness via the pentose phosphate pathway. These different pathways of NADPH
production enable the regulation of diverse metabolic pathways in chloroplasts by the
NADPH-dependent thioredoxin system.
In this thesis, the role of NADPH-dependent thioredoxin system in the redox-control of
chloroplast development and metabolism was studied by characterization of
Arabidopsis thaliana T-DNA insertion lines of NTRC gene (ntrc) and by identification
of chloroplast proteins regulated by NTRC. The ntrc plants showed the strongest
visible phenotypes when grown under short 8-h photoperiod. This indicates that i)
chloroplast NADPH-dependent thioredoxin system is non-redundant to ferredoxinthioredoxin system and that ii) NTRC particularly controls the chloroplast processes
that are easily imbalanced in daily light/dark rhythms with short day and long night. I
identified four processes and the redox-regulated proteins therein that are potentially
regulated by NTRC; i) chloroplast development, ii) starch biosynthesis, iii) aromatic
amino acid biosynthesis and iv) detoxification of H2O2. Such regulation can be
achieved directly by modulating the redox state of intramolecular or intermolecular
disulfide bridges of enzymes, or by protecting enzymes from oxidation in conjunction
with 2-cysteine peroxiredoxins. This thesis work also demonstrated that the enzymatic
antioxidant systems in chloroplasts, ascorbate peroxidases, superoxide dismutase and
NTRC-dependent 2-cysteine peroxiredoxins are tightly linked up to prevent the
detrimental accumulation of reactive oxygen species in plants.
TABLE OF CONTENTS
LIST OF ORIGINAL PUBLICATIONS
ABBREVIATIONS
ABSTRACT
1. INTRODUCTION..................................................................................................................... 9
1.1. CHLOROPLASTS IN PLANTS................................................................................................... 9
1.2. PHOTOSYNTHESIS AND OTHER METABOLIC REACTIONS IN CHLOROPLASTS ....................... 10
1.2.1. Photosynthetic light reactions ................................................................................... 10
1.2.2. Calvin-Benson cycle and other metabolic pathways in chloroplasts ....................... 11
1.3. REGULATORY AND ANTIOXIDANT REDOX-SYSTEMS IN CHLOROPLAST .............................. 13
1.3.1. Thioredoxin systems .................................................................................................. 14
1.3.2. Reactive oxygen species in chloroplast ..................................................................... 17
1.3.3. Detoxification of reactive oxygen species ................................................................. 18
1.4. CHLOROPLAST BIOGENESIS ................................................................................................ 20
1.4.1. Chlorophyll biosynthesis ........................................................................................... 22
1.5. REGULATION OF CHLOROPLAST BIOGENESIS AND ACCLIMATION TO ENVIRONMENTAL
CUES .......................................................................................................................................... 24
1.5.1. Anterograde signaling ............................................................................................... 25
1.5.2. Chloroplast-to-nucleus retrograde signaling ........................................................... 25
1.5.3. ROS in plant signaling .............................................................................................. 27
2. AIMS OF THE STUDY.......................................................................................................... 28
3. METHODOLOGICAL ASPECTS ....................................................................................... 29
3.1. PLANT MATERIAL AND GROWTH CONDITIONS .................................................................... 29
3.2. STRESS TREATMENTS ......................................................................................................... 29
3.3. ANALYSIS OF PIGMENTS AND PROTEINS ............................................................................. 30
3.4. MICROARRAY ANALYSIS .................................................................................................... 30
3.5. BIOPHYSICAL METHODS ..................................................................................................... 30
3.6. MICROSCOPY ..................................................................................................................... 31
3.7. IN VIVO-DETECTION OF H2O2 AND SUPEROXIDE................................................................. 31
3.8. ANALYSES OF AMINO ACIDS, HORMONES AND SUGARS...................................................... 31
3.9. YEAST TWO-HYBRID ANALYSIS .......................................................................................... 32
4. OVERVIEW OF THE RESULTS......................................................................................... 33
4.1. DAILY LIGHT/DARK RHYTHM MODULATES THE PHOTOSYNTHESIS AND GROWTH OF
ARABIDOPSIS ............................................................................................................................ 33
4.1.1. Impact of photoperiod on growth and photosynthesis in wild type Arabidopsis ...... 33
4.1.2. Impact of photoperiod on the redox metabolism of mesophyll cells ......................... 34
4.1.3. Knockout of NTRC impairs the acclimation of Arabidopsis to short photoperiods . 35
4.1.4. Photosynthesis of knockout ntrc lines ....................................................................... 35
4.2. IDENTIFICATION OF CHLOROPLAST PROCESSES CONTROLLED BY NADPH-DEPENDENT
THIOREDOXIN SYSTEM .............................................................................................................. 36
4.2.1. Impact of NTRC on chloroplast biogenesis .............................................................. 36
4.2.2. Starch and sucrose metabolism in knockout ntrc leaves........................................... 38
4.2.3. NTRC and chloroplast ROS metabolism ................................................................... 39
4.2.4. Knockout of NTRC interferes with the homeostasis of metabolites produced in
shikimate pathway ............................................................................................................... 39
4.3. CONTRIBUTION OF CHLOROPLAST ASCORBATE PEROXIDASES TO THE ROS METABOLISM IN
ARABIDOPSIS ............................................................................................................................ 41
5. DISCUSSION .......................................................................................................................... 42
5.1. NTRC IN REDOX-REGULATION OF CHLOROPLAST BIOGENESIS .......................................... 42
5.1.1. Regulation of chloroplast division by NTRC ............................................................ 43
5.1.2. Regulation of chlorophyll biosynthesis by NTRC ..................................................... 44
5.1.3. Thylakoid formation in ntrc mutant lines .................................................................. 45
5.2. REGULATION OF CHLOROPLAST METABOLISM BY NTRC .................................................. 45
5.2.1. Redox control of starch metabolism in chloroplasts ................................................. 45
5.2.2. Altered homeostasis of metabolites synthesized from the end products of shikimate
acid pathway........................................................................................................................ 46
5.3. THIOREDOXIN- AND ASCORBATE-DEPENDENT SCAVENGING OF ROS IN CHLOROPLASTS .. 47
5.3.1. Detoxification of H2O2 by NTRC ............................................................................... 47
5.3.2. Impacts of chloroplast APXs on chloroplast redox homeostasis .............................. 48
5.4. PHOTOPERIODIC DEVELOPMENT OF PHOTOSYNTHETIC TISSUES ......................................... 49
5.4.1. Impact of NTRC knockout on the growth of Arabidopsis ......................................... 50
5.5. CHLOROPLAST-TO-NUCLEUS RETROGRADE SIGNALING ..................................................... 50
6. CONCLUDING REMARKS ................................................................................................. 52
ACKNOWLEDGEMENTS ....................................................................................................... 53
REFERENCES ............................................................................................................................ 54
ORIGINAL PUBLICATIONS .................................................................................................. 63
“There is a theory which states that if ever anyone discovers
exactly what the Universe is for and why it is here,
it will instantly disappear and be replaced by something
even more bizarre and inexplicable.
There is another theory which states that this has already happened.”
Douglas Adams: The Restaurant at the End of the Universe (1980)
Introduction
9
1. Introduction
Chloroplasts arose approximately 1.5 billion years ago when a free-living autotrophic
bacterium, an ancestor of present-day cyanobacterium was engulfed by a
mitochondrion-containing eukayotic host cell and became an endosymbiont. This event
provided mutual benefit for both partners; bacterium occupied an untapped niche and
the host received nutrients such as reduced carbon (Glynn, et al., 2007). Since its
origin, the chloroplast has become fully integrated to the life cycle of photosynthetic
eukaryotes, being essential to the plants themselves but also enabling the existence of
heterotrophic life on Earth.
1.1. Chloroplasts in plants
Chloroplasts (Figure 1) are flat disc-shaped organelles, surrounded by an envelope that
consists of an outer and inner lipid membrane. Between these two layers is the
intermembrane space. Inside the chloroplast, the third membrane system, thylakoids,
appear individually as stroma thylakoids or form stacked grana structures. Thylakoids
of a single chloroplast form a three-dimensional interconnected network, enclosing a
space called lumen (Mustardy and Garab, 2003). The chloroplast content outside the
thylakoids, the stroma, corresponds to the cytosol of the original endosymbiotic
bacterium, containing one or more molecules of circular DNA as well as ribosomes,
proteins, metabolites and temporary starch granules. A typical Arabidopsis thaliana
(Arabidopsis) leaf cell contains over 100 chloroplasts.
Figure 1. Outline of the
chloroplast ultrastructure. 1. outer
membrane, 2. intermembrane
space, 3. inner membrane
(1+2+3 : envelope), 4. stroma, 5.
thylakoid membrane 6. thylakoid
lumen (inside of thylakoid), 7.
stroma thylakoid, 8. granum
(stack of thylakoids)
The size of plastidial DNA (plastome) is significantly smaller than the cyanobacterial
genome; chloroplast genome encodes 50-200 proteins (Martin, 2003), whereas the
genome of a cyanobacterium Synechocystis theoretically encodes more than 3600
proteins (Battchikova, et al., 2010). During the course of evolution, genes have been
transferred from chloroplast to nucleus where they can be regulated in an integrated
10
Introduction
manner (Martin and Herrmann, 1998). Some genes are still retained in chloroplasts,
most probably because the expression of those genes is required to be directly and
efficiently regulated by the factors present in chloroplast (Allen, et al., 2005).
1.2. Photosynthesis and other metabolic reactions in chloroplasts
The chloroplasts’ ability to perform photosynthesis, the conversion of carbon dioxide
into organic compounds using the energy of sunlight, is vital for life on Earth. Sugars
produced by photosynthesis provide nourishment, either directly or indirectly, to nearly
all life forms and the oxygen released in the process as a side product maintains the
oxygen level in the atmosphere and enables the living of aerobic life forms.
Photosynthesis consists of two distinct phases that both occur inside the chloroplast,
yet in different compartments. The light-dependent reactions take place in thylakoid
membranes, in which light energy drives electron flow between a series of multisubunit protein complexes, eventually generating ATP and reducing equivalents. This
chemical energy is then spent in the stroma by Calvin-Benson cycle to fix CO2 into
energy-rich sugar phosphates. Sugar phosphates are then exported to the cytosol or
stored in the chloroplast as starch.
1.2.1. Photosynthetic light reactions
In plants, the photosynthetic light-induced electron transfer reactions are carried out by
two physically separated, thylakoid-membrane-bound multi-subunit protein complexes,
photosystem I and II (PSI and PSII) in addition to cytochrome b6f (Cyt b6f) complex,
ATP synthase and mobile electron carriers. The electron transfer reactions begin when
a photon of visible light is absorbed by the reaction center chlorophyll molecule in PSII
and in PSI (Figure 2). The reaction center pigments of photosystems are able to absorb
photons directly, but light is also absorbed by the light-harvesting complex, a set of
photosynthetic pigments outside the reaction centers of each photosystem. Lightharvesting complex functions as a solar collector that feeds light energy through
resonance energy transfer to the reaction center chlorophyll molecules at photosystems.
The absorbed energy boosts the PSII reaction center pigment P680 to an excited state
P680* which rapidly transfers an electron to a nearby pheophytin, a PSII-bound
pigment. The electron flows from pheophytin via PSII-bound plastoquinones QA and
QB to plastoquinone (PQ) pool, mobile electron carriers that pass electrons to Cyt b6f
complex. When P680* delivers an electron, P680+ is formed. It is a very strong oxidant
capable of extracting electrons from water molecules bound at the manganese cluster in
oxygen evolving complex (OEC), which results in splitting water to protons and
oxygen.
Introduction
11
Figure 2. Schematic diagram of linear electron transfer reactions in oxygenic photosynthesis.
See text for details.
Electrons from the Cyt b6f complex are transferred to PSI by plastocyanin (PC),
another mobile electron carrier. Before an electron can be accepted by PSI reaction
center pigment P700, another photon of light is needed to excite P700 to P700*. When
P700* releases its electron to the primary electron acceptor of PSI, P700+ accepts the
electron delivered by PC and returns to the ground state. The electrons are finally
accepted by ferredoxin (Fd) which then reduces NADP+ to NADPH with the help of
ferredoxin-NADP+ oxidoreductase (FNR). Both the proton translocation coupled to
electron transfer and the release of protons by the water oxidation reaction contribute
to the electrochemical gradient across the thylakoid membrane that drives the synthesis
of ATP by ATP synthase. ATP and NADPH are used in the Calvin-Benson cycle in the
stroma to fix CO2 to carbohydrates.
1.2.2. Calvin-Benson cycle and other metabolic pathways in chloroplasts
The reactions in Calvin-Benson cycle catalyze the fixation of carbon dioxide
and the reduction of sugars in three stages: (i) the incorporation of a CO2
molecule into ribulose 1,5-bisphosphate to form two molecules of 3phosphoglycerate (PGA) in the reaction catalyzed by Rubisco; (ii) the reduction
of PGA to glyceraldehyde 3-phosphate (G3P) that is used to form hexose
sugars; and (iii) the transformation of five of the six G3P into three ribulose
1,5-bisphosphates to replace the ones that initiated the cycle (Figure 3).
12
Introduction
Figure 3. Carbon assimilated via Calvin-Benson cycle is partitioned into fraction retained in
chloroplast for starch synthesis and a fraction exported to the cytosol for sucrose synthesis.
RuBP, ribulose 1,5-bisphosphate; PGA, 3-phosphoglycerate; G3P, glyceraldehyde 3-phosphate;
Fru6P, fructose 6-phosphate; Glc6P, glucose 6-phosphate; Glc1P, glucose 1-phosphate;
AGPase, ADP-glucose pyrophosphorylase; ADPGlc, ADP-glucose; PPi, pyrophosphate; Pi
inorganic phosphate; UDPGlc, UDP-glucose; TP/Pi translocator, triose phosphate/Pi
translocator.
Photoassimilated carbon, in the form of G3P, is converted into hexose phosphates that
serve as precursors for the synthesis of starch and sucrose. Starch is synthesized within
the stroma of chloroplasts (recently reviewed by Zeeman, et al., 2007 and 2010). In the
key step of starch synthesis, glucose 1-phosphate is activated with ATP in a reaction
catalyzed by ADP-glucose pyrophosphorylase (AGPase) to yield ADP-glucose and
pyrophosphate that is hydrolyzed to two inorganic phosphate groups (Pi) by alkaline
pyrophosphatase. ADP-glucose then transfers the glucosyl group to the nonreducing
end of a starch molecule (Figure 3). The Pi released in the synthesis of starch operates
as a regulatory link between light reactions and carbon reactions since it is consumed
in the synthesis of ATP in the light reactions. Starch is the major storage form of
chemical energy in most plants. It is transiently stored in chloroplasts and degraded
during dark periods to provide sugars for metabolism and growth throughout the night.
Starch degradation begins with the reversible phosphorylation of starch granules by
glucan, water dikinase (GWD) and phosphoglucan phosphatase (DSP4). Soluble
Introduction
13
glucans are then released from the starch granule by amylases and converted to glucose
and maltose that are exported from the chloroplast to the cytosol (Zeeman, et al.,
2010).
Sucrose is the transport form of carbohydrates in plants and in contrast to starch,
sucrose is synthesized in the cytosol. G3P is transported to cytosol via triose
phosphate/Pi translocator that regulates the competing processes, the synthesis of starch
and sucrose in mesophyll cells. When the cytosolic Pi concentration is high, G3P is
exported to cytosol and converted to sucrose. When the cytosolic Pi concentration is
low, starch synthesis is activated inside the chloroplasts (Figure 3).
In the sucrose synthesis glucose 1-phosphate is activated with UTP to form UDPglucose, which then transfers a glucosyl group to fructose 6-phosphate to form sucrose
6-phosphate. Hydrolysis of sucrose 6-phosphate by sucrose phosphatase yields sucrose
(Figure 3).
Besides photosynthesis a number of other metabolic processes are localized to
chloroplasts in plants. Nitrogen assimilation, the biosynthesis of fatty acids, amino
acids, vitamins, purines, hormones and heme take place in chloroplasts (Galili, 2002;
Weber and Flugge, 2002; Boldt and Zrenner, 2003; Lunn, 2007; Tanaka and Tanaka,
2006). These pathways are not necessarily confined exclusively to chloroplasts as part
of them is also, partially or alternatively conducted in the cytosol and in the
mitochondria. Thus the pathways in different cellular compartments need to be linked
together, which is usually attained by integral membrane translocator proteins that
transfer intermediate metabolites. The transported metabolites also act as signals that
provide information of metabolic conditions in the originating compartment (Lunn,
2007).
1.3. Regulatory and antioxidant redox-systems in chloroplast
Chloroplasts contain numerous redox-active components that function in electron
transfer reactions, as oxidants or as specific regulators of other molecules. The redox
compounds in chloroplasts may also participate in signaling cascades that control
metabolic reactions, gene transcription and translation not only in the chloroplast but
via retrograde signaling in the nucleus as well. The redox-active compounds in
chloroplasts include photosynthetic electron transport chain components in thylakoid
membranes and electron-flow-dependent soluble components such as ROS,
thioredoxins and glutaredoxins as well as antioxidants. In this chapter the redox
compounds related to this thesis work, thioredoxins, ROS and antioxidative enzymes,
are introduced.
14
Introduction
1.3.1. Thioredoxin systems
Thioredoxins (Trxs) are small proteins with the conserved amino acid sequence of
WCG/PPC that has thiol:disulfide oxidoreductase activity. These regulatory proteins
have a low redox potential (between -285 and -350 mV) that gives them strong
reductive properties (Gelhaye, et al., 2005). In the reduced state, Trxs are able to
reduce the disulfide bridges formed between redox-active cysteines in the Trx target
proteins. Subsequently, the oxidized Trxs are reduced by thioredoxin reductases,
forming together the so-called thioredoxin system. In contrast to non-photosynthetic
organisms, plants contain a particularly large number of Trx isoforms in several
cellular compartments, including the cytosol, nucleus, mitochondria and chloroplasts.
In Arabidopsis, 42 genes coding for Trx and Trx-like sequences have been identified
(Table 1; Meyer, et al., 2008; Chibani, et al., 2009; Arsova, et al., 2010), and many of
these proteins are located in the chloroplasts or at least predicted to be imported into
the chloroplasts. Chloroplasts contain f-, m-, x-, y-, z- type Trxs and a drought-induced
stress protein (CDSP)32; h-type Trxs are present both in cytosol and mitochondria, and
o-type Trxs are present only in mitochondria. The crystal structures of f-, m- and h-type
Trxs and their target proteins have been resolved (Chiadmi, et al., 1999; Johansson, et
al., 1999; Capitani, et al., 2000; Fermani, et al., 2001; Falini, et al., 2003; Coudevylle,
et al., 2005; Peterson, et al., 2005; Maeda, et al., 2010) whereas even the function of
other Trxs, especially that of the Trx-like proteins is yet fairly unknown. The reasons
for the high number of Trxs in plants compared to other living organisms are still
unclear. It has been suggested to be related to the increased generation of ROS during
photosynthesis and also to genome duplications in terrestrial plants (Meyer, et al.,
2008).
Oxidized Trxs become reduced via the action of thioredoxin reductases. Cytosolic and
mitochondrial Trxs in plants are reduced by NADPH-dependent thioredoxin reductases
(NTRs) whereas the reduction of Trxs in chloroplasts is accomplished via the action of
two systems, NADPH-dependent thioredoxin reductase C (NTRC) and ferredoxinthioredoxin reductase (FTR). Arabidopsis has three genes encoding the isoforms of
NTRs. Two isoforms, NTRA and NTRB are localized both in cytosol and
mitochondria and constitute the Trx-reduction system in those cellular compartments
(Serrato, et al., 2004). NTRC is a 55-kDa fusion protein localized to chloroplasts
(Serrato, et al., 2004). It consists of two domains, a thioredoxin reductase domain in
the N-terminus and a Trx domain in the C-terminus (Figure 4, Serrato, et al., 2004) that
is missing from other NTR isoforms (Serrato, et al., 2004). Both NTRC domains
possess a redox-active cysteine pair.
Introduction
15
Table 1. Thioredoxins in Arabidopsis. From Meyer et al. (2008).
Thioredoxin type
AGI ID
Common name
F
AT3G02730
AT5G16400
AT1G03680
AT4G03520
AT2G15570
AT3G15360
AT1G50320
AT1G76760
AT1G43560
AT3G06730
AT1G76080
AT4G37200
AT5G06690
AT5G04260
AT1G08570
AT4G29670
AT5G61440
AT2G33270
AT1G07700
AT2G35010
AT1G31020
AT3G51030
AT5G42980
AT1G19730
AT1G45145
AT5G39950
AT1G59730
AT1G69880
AT2G40790
AT3G08710
AT1G11530
AT3G56420
AT3G53220
AT3G06730
AT5G42850
AT3G17880
AT1G60420
AT4G31240
AT4G04950
AT4G32580
AT1G52990
AT2G41680
f1
f2
m1
m2
m3
m4
x
y1
y2
z
CDSP32
HCF164
WCRKC1
WCRKC2
Lilium1
Lilium2
Lilium3
Lilium4
Lilium5
o1
o2
h1
h3
h4
h5
h2
h7
h8
CxxS1
h9
CxxS2
CxxC2
WCGVC
CI
Clot
TDX
Nucleor1
Nucleor2
Picot1
Picot2
TARWCGPC
NTRC
M
X
Y
Z
CDSP32
HCF164
WCRKC
Lilium
O
h subtype I
h subtype II
h subtype III
WCGVC
Cf-6 Interacting
Clot TRP14
TDX
Nucleoredoxin
Picot
TARWCGPC
NTRC
Subcellular localization in
Arabidopsis
Chloroplast
Chloroplast
Chloroplast
Chloroplast
Chloroplast
Chloroplast
Chloroplast
Chloroplast
Chloroplast
Chloroplast
Chloroplast
Chloroplast
Chloroplast (homology)
Chloroplast (homology)
Chloroplast (homology)
Chloroplast (homology)
Chloroplast (homology)
Chloroplast (homology)
Chloroplast (homology)
Mitochondria
?
Cytosol
Cytosol
Cytosol
Cytosol
Cytosol
Cytosol (homology)
Cytosol (homology)
Cytosol (homology)
Cytosol (homology)
Cytosol (homology)
Cytosol (homology)
Cytosol (homology)
?
Cytosol (homology)
Cytosol/nucleus
Nucleus (homology)
Nucleus (homology)
Cytosol (homology)
Cytosol (homology)
Secretion (homology)
Chloroplast
16
Introduction
Figure 4. Schematic model of NTRC domains and the reaction mechanism. NTR, NADPHdependent thioredoxin reductase domain containing binding sites for NADPH and FAD; Trx,
thioredoxin domain; CAIC and CGPS, redox-active sites of NTR domain and Trx domain,
respectively. Arrows represent electron transfer routes; see text for details.
Several NTR crystal structures have been resolved; most of them are bacterial enzymes
(Kuriyan, et al., 1991; Lennon, et al., 1999 and 2000; Waksman, et al., 1994; Akif, et
al., 2005; Hernandez, et al., 2008; Ruggiero, et al., 2009), but also two structures for
plant enzymes, Arabidopsis NTRB (Dai, et al., 1996) and HvNTR2 from barley
(Kirkensgaard, et al., 2009) have been published. Based on those structures, a model
for NTRC reaction mechanism has been proposed (Perez-Ruiz and Cejudo, 2009).
According to this model, NTRC functions as a homodimer. Both subunits are able to
transfer electrons from NADPH to FAD bound to NTR domain, and from FAD to the
adjacent redox-active disulphide in the NTR domain (Figure 4). From the NTR domain
of one subunit, electrons are then transferred via inter-subunit reaction to the redoxactive site of Trx domain of the other subunit. The Trx domain is then capable of
reducing its target proteins (Figure 4). As NADPH can also be produced during the
dark period via the action of pentose phosphate pathway (Neuhaus and Emes, 2000), it
is suggested that NADPH-dependent reduction of thioredoxins play an important role
in plant chloroplasts during the night or under low light, when the level of reduced
ferredoxin is low (Perez-Ruiz, et al., 2006).
Another plastidial thioredoxin reductase, FTR has been intensively studied since its
discovery in the 1970’s (Wolosiuk and Buchanan, 1978). FTR uses ferredoxin, reduced
by electrons transported through the photosystems during photosynthesis, as an
electron donor, thereby converting a light-activated electron signal to thiol signal. FTR
in higher plants is an αβ-heterodimer encoded by the nuclear genes. FTR is comprised
of a 13-kDa large catalytic subunit containing both a [4Fe-4S] cluster and a proximal
redox-active disulfide, and a 8-12 kDa variable subunit (Schürmann and Jacquot, 2000;
Schürmann and Buchanan, 2008). FTR is a versatile enzyme regarding its reactivity as
it efficiently reduces f- and m-type Trxs (Schürmann and Buchanan, 2008). Specificity
arises from the interaction between Trxs and target enzyme. Trx f functions primarily
Introduction
17
in the activation of enzymes in photosynthetic metabolism like Calvin-Benson cycle
enzymes (reviewed by Schürmann and Buchanan, 2008), Rubisco activase (Zhang and
Portis, 1999) and ATP synthase (Schwarz, et al., 1997; Stumpp, et al., 1999). These
enzymes interact less efficiently with Trx m. NADP-malate dehydrogenase was
originally considered to be specifically activated by Trx m, but was later shown to be
even more efficiently activated by Trx f (Hodges, et al., 1994; Geck, et al., 1996). Both
Trx f and Trx m are also active in enzyme deactivation (Schürmann and Buchanan,
2008). For example, the STN7 protein kinase that phosphorylates thylakoid lightharvesting proteins loses its activity after treatment with Trx f and Trx m (Rintamäki, et
al., 2000).
The development of proteomic approaches during recent years has allowed the
identification of numerous proteins linked to thioredoxins. In addition to
photosynthesis, thioredoxins appear to play important roles in a number of
physiological processes in plants, including housekeeping metabolism, development
and abiotic and biotic stress defense mechanisms (Montrichard, et al., 2009). However,
although thioredoxins are well characterized in the control of photosynthesis (reviewed
by Schürmann and Buchanan, 2008), the functions of many newly identified
thioredoxin target proteins still remain to be explored.
1.3.2. Reactive oxygen species in chloroplast
Photosynthesis produces intermediates with extreme redox potentials: PSI generates a
strong reductant capable of reducing NADP+ and PSII reduces the electron donors to
PSI and generates a very strong oxidant capable of oxidizing water to molecular
oxygen. These basic photosynthetic reactions pose a significant risk for electron
transfer to oxygen in chloroplast. Such reactions are harmful to the organism since
oxygen can be considered as a double-edged molecular sward as it plays two different
roles in biological systems; it is vital for aerobic metabolism but at the same time the
reduction of molecular oxygen may result in formation of reactive oxygen species
(ROS) that can harm the constituents of the cells and ultimately even cause a cell
death.
The PSI and PSII complexes in thylakoid membranes are indeed the major generation
sites of ROS in chloroplasts. Photoreduction of oxygen to hydrogen peroxide (H2O2) in
PSI was discovered over 50 years ago by (Mehler, 1951). The primary step in this
reaction generates superoxide radical (O2.-), and its spontaneous or enzymatic
disproportionation produces hydrogen peroxide (H2O2) and O2 (Figure 5; Asada, et al.,
1974). On the other hand, in PSII, oxygen of the ground (triplet) state (3O2) may be
excited to singlet state (1O2) by the triplet state of excited chlorophyll (Telfer, et al.,
1994; Hideg, et al., 1998; Krieger-Liszkay, 2005). The production of ROS in light is
largely affected by physiological and environmental factors; the rate is enhanced under
conditions where electron transport chain is overreduced, that is when light absorbed
exceeds the dissipation and utilization capacity of light energy in photosynthesis (Li, et
18
Introduction
al., 2009). Due to the high reactivity of ROS they pose a continuous threat to cellular
constituents for uncontrolled oxidation of lipids, nucleic acids and amino acids.
Figure 5. Generation of different forms of ROS from oxygen. See text for details. Figure drawn
according to Gill and Tuteja (2010).
H2O2 is a moderately reactive form of oxygen. It is not a free radical and therefore less
reactive and longer-lived compound than superoxide anion. On the other hand, due to
its lower reactivity compared to other ROS, H2O2 can diffuse out of the chloroplasts
and within and between cells and act beyond the generation site (Mubarakshina, et al.,
2010). In principle, H2O2 is relatively stable in the absence of transition metals.
However, with molecules containing Fe2+ or other transition metals, hydroxyl radicals
can be formed from H2O2 via Fenton reaction (Figure 5; reviewed by Blokhina et al.,
2003). Hydroxyl radical is highly toxic ROS that can potentially react with all
important biological molecules including DNA, proteins and lipids (Gill and Tuteja,
2010).
1.3.3. Detoxification of reactive oxygen species
To minimize the potentially hazardous reactions initiated by ROS, plants have evolved
an extensive antioxidant defence system with low-molecular mass non-enzymatic
antioxidants such as ascorbate, glutathione, tocopherols, carotenoids and phenolic
compounds, and antioxidant enzymes that respond to redox imbalances (Mittler, et al.,
2004). Under physiological steady state conditions plants typically maintain ROS at
low levels, but in the induction of photo-oxidative stress both the production of ROS
and the pool sizes of low-molecular mass antioxidants and antioxidant enzymes
increase (Foyer, et al., 1997). In contrast to most organisms, plants have multiple
genes encoding for antioxidant enzymes. Different isoforms are specifically targeted to
Introduction
19
chloroplasts, mitochondria, peroxisomes, as well as to the cytosol and apoplast (Apel
and Hirt, 2004; Mittler, et al., 2004).
Detoxification of superoxide anions and H2O2 in chloroplasts is catalyzed by four
antioxidant enzyme systems presented in Figure 6. Superoxide anions produced by PSI
are rapidly dismutated to H2O2 either spontaneously or by superoxide dismutases
(SODs) (Foyer et al 1997; Asada, 1999) (Figure 6A). Subsequently, H2O2 is converted
to water by ascorbate peroxidases (APXs) and ascorbate, by peroxiredoxins (Prxs) and
thioredoxins, or by glutathione peroxidase (GPX) cycle (Foyer, et al., 1997; Asada,
1999; Mittler, 2002; Rouhier and Jacquot, 2002).
Figure 6. Overview of the enzymatic ROS scavenging reactions by SODs (A), APXs (B), Prxs
(C) and GPXs (D) in chloroplasts. See text for details.
APXs are haem-binding enzymes that reduce H2O2 with ascorbate as an electron donor.
Plants contain five APX isoenzymes in different cellular compartments. Two
chloroplastic isoforms, a 38 kDa thylakoid-bound APX (tAPX) and a 33 kDa stromal
APX (sAPX), which is dually targeted also to mitochondria, are both encoded by a
20
Introduction
single nuclear gene in Arabidopsis. 25 kDa APX1 and APX2 are located in the
cytoplasm and a 31 kDa APX3 has been found in peroxisomes and oilseed
glyoxysomes. In the first APX-catalyzed step of ascorbate-glutahione cycle (Figure
6B), H2O2 is reduced to water with electrons from ascorbate which is oxidized into
monodehydroascorbate (MDA). In the second step, MDA reductase (MDAR) reduces
MDA into ascorbate with the help of NAD(P)H. MDA can also become oxidized
spontaneously to dehydroascorbate (DHA) that is reduced to ascorbate by DHA
reductase (DHAR) or by glutaredoxins with the help of GSH that is oxidized to GSSG.
The cycle closes with glutathione reductase (GR) that rereduces GSSG back into GSH
with NADPH (Asada, 1999).
Prxs form a large group of 17-22 kDa enzymes which contain a conserved cysteine in
the N-terminus of a protein that is responsible for peroxidase activity (Rouhier and
Jacquot, 2002). In Arabidopsis, four Prxs are found in chloroplasts, 2-cysteine (2-Cys)
Prxs A and B, PrxQ and PrxIIE. Like APXs, Prxs rely on external electron donor in
order to reduce H2O2 to water (Figure 6C). A regular electron donor for chloroplastic
Prx isoforms is a reduced thioredoxin (Rouhier and Jacquot, 2002), which is oxidized
in the reaction and then needs to be re-reduced by thioredoxin reductase.
The GPX cycle converts H2O2 into water using reducing equivalents from glutathione
(GSH) (Figure 6D) (Ursini, et al., 1995) or thioredoxins (Herbette, et al., 2002; Jung, et
al., 2002). Oxidized GSSG is again converted into GSH by glutathione reductase with
NADPH as a reducing agent.
1.4. Chloroplast biogenesis
Chloroplast biogenesis in higher plants is defined here as a process in which
chloroplasts develop from proplastids, small precursor organelles that lack organized
internal membranes and are primarily present in meristematic cells. As meristematic
cells differentiate to leaf mesophyll cells, proplastids differentiate into chloroplasts
(Vothknecht and Westhoff, 2001). In young, developing leaves the conversion of
proplastids to chloroplasts can occur directly or indirectly via etioplast and etiochloroplast stages (Figure 7). Direct transformation of proplastids to chloroplasts
requires high light intensity, while etioplasts and etio-chloroplasts are developed under
darkness and also under low light and natural light-dark cycles (Solymosi and Schoefs,
2010). In contrast to proplastids, etioplasts contain an inner membrane system which is
organized in highly regular paracrystalline structures called prolamellar bodies (PLBs),
several proteins involved in photosynthesis (Blomqvist, et al., 2008; Kanervo, et al.,
2008) and protochlorophyllide, a precursor for chlorophyll (Schoefs and Franck, 2003).
Under natural light-dark cycles PLBs appear during dark periods and disappear in the
beginning of the light period (Solymosi and Schoefs, 2010). PLB membranes are
precursors for thylakoid membranes that are formed during greening when etioplasts
are converted to chloroplasts.
Introduction
21
During the conversion of proplastids into chloroplasts, the plastid volume increases up
to 100-fold (Gutierrez-Nava, et al., 2004). Plastids also need to increase their number
by binary fission as their ‘host’ cells expand. The total volume of chloroplasts in cell
and the size of mesophyll cell are strictly controlled in plant species (Pyke, 1999) but
the regulation mechanisms are still fairly unknown. Plastid division requires the
operation of several plastid division proteins (recently reviewed by Miyagishima and
Kabeya, 2010). FtsZ proteins self-assemble beneath the inner envelope membrane to
form the core ring structure (Z-ring) on which other division components assemble.
The correct position of the Z-ring in the middle of the chloroplast is determined by a
complex of Min proteins and ARC3 which is localized in the ends of chloroplast thus
preventing the assembly of the Z-ring at incorrect position (Maple and Møller, 2007).
Figure 7. Different pathways for chloroplast development. Under low light intensities and
under natural light-dark cycles, proplastids differentiate to etio-chloroplasts (1.) before
conversion to chloroplasts (2.). This route is reversible during the dark periods until
chloroplasts are fully maturated (3.). The proplastids of seedlings germinating in the absence of
light differentiate into etioplasts (4.). When provided with light, etioplasts are transformed into
chloroplasts via the etio-chloroplast stage (5. and 2.). Under high light intensities, proplastids
differentiate directly to chloroplasts (6.). Figure drawn according to Solymosi and Schoefs
(2010).
22
Introduction
Division and expansion of chloroplasts also require active synthesis of membranes in
chloroplasts. New thylakoid membranes are formed from lipids that are either derived
from precursors assembled de novo in the plastid or imported from the endoplasmic
reticulum (Benning, 2009). Thylakoid membranes are constructed via an orchestrated
process in which membrane lipids, proteins and pigments are assembled into
functional, three-dimensional structures (Vothknecht and Westhoff, 2001). Since
relatively small number of genes resides in chloroplasts, the majority of plastid
proteins are encoded in the nucleus, translated in the cytosol and imported into
chloroplasts. Chloroplast multiprotein complexes such as ribosomes and photosystems
are patchworks of subunits encoded by both chloroplast and nuclear genomes. In order
to attain functional thylakoid membranes, biosynthesis and import of these components
must be well-balanced. Mutations that interfere with the synthesis of major protein
components of thylakoid membranes, e.g. major subunits of photosystems (Meurer, et
al., 1998; Baena-Gonzalez, et al., 2003; Ido, et al., 2009), protein import apparatus
(Bauer, et al., 2000) or pigment biosynthesis (Falbel and Staehelin, 1994; Masuda, et
al., 2003) may cause defects in thylakoid formation. Furthermore, specific chloroplast
proteins involved in thylakoid formation have been recently identified. THYLAKOID
FORMATION 1 (THF1) was shown to control an important step required for the
normal organization of membrane vesicles into mature thylakoid stacks (Gao, et al.,
2006), and loss of FZO-LIKE (FZL) protein function resulted in altered chloroplast
and thylakoid morphology in Arabidopsis (Wang, et al., 2004). Also VESICLE
INDUCING PLASTID PROTEIN 1 (VIPP1) has been suggested to be important for
thylakoid membrane formation, since mutant plants with greatly reduced expression of
vipp1 gene show inhibited chloroplast vesicle transport and probably therefore have
lost the ability to build up a proper thylakoid membrane system (Kroll, et al., 2001;
Aseeva, et al., 2007).
1.4.1. Chlorophyll biosynthesis
Chlorophyll, the green pigment, is vital for photosynthesis as it absorbs light energy
and directs the light energy towards photosystems. In plants, chlorophyll biosynthesis
takes place in chloroplasts, and all participating enzymes are encoded by the nucleus.
Chlorophyll biosynthesis (reviewed by Mochizuki et al., 2010) starts with the
formation of 5-aminolevulinic acid (ALA) from glutamyl-tRNAGlu by glutamyl-tRNA
reductase (HEMA1) and glutamate 1-semialdehyde aminotransferase (Figure 8). Eight
molecules of ALA are then assembled via three enzymatic steps into uroporphyrinogen
III, which can continue in the chlorophyll biosynthesis pathway or be converted to
siroheme, the prosthetic group for both nitrite and sulfite reductases in plants. In the
subsequent step of chlorophyll biosynthesis, uroporphyrinogen III is decarboxylated to
form protoporphyrinogen IX, which is further oxidized to protoporphyrin IX.
Protoporphyrin IX can be converted to heme via insertion of Fe2+ or to chlorophyll via
insertion of Mg2+, thus being the second branchpoint of the pathway. In the chlorophyll
branch, insertion of Mg2+ is catalyzed by Mg-chelatase, a multi-subunit protein
complex. Mg-protoporphyrin IX is then modified to Mg-protoporphyrin monomethyl
Introduction
23
ester by Mg-protoporphyrin IX methyltransferase (CHLM), and in subsequent step
catalyzed by Mg-protoporphyrin monomethylester cyclase (CRD1) to form
protochlorophyllide which is then reduced to chlorophyllide a. The reaction of
protochlorophyllide
reduction
to
chlorophyllide,
mediated
by
NADPH:protochlorophyllide oxidoreductase (POR), is catalyzed by the energy of
light. Chlorophyllide a is esterified with a phytol chain to give chlorophyll a, some of
which is reversibly converted to chlorophyll b.
Figure 8. Schematic illustration of the chlorophyll biosynthesis pathway. HEMA, glutamyltRNA reductase; GSA, glutamate-1-semialdehyde 2,1-aminomutase; ALAD5-aminolevulinate
dehydratase; PBGD, porphobilinogen deaminase; UROS, uroporphyrinogen III synthase;
UROD, uroporphyrinogen III decarboxylase; CPO, coproporphyrinogen III oxidase; PPO,
protoporphyrinogen IX oxidase; GUN4, regulator of Mg-chelatase; CHLD, Mg-chelatase Dsubunit; CHLI, Mg-chelatase I-subunit; CHLH, Mg-chelatase H-subunit; CHLM, Mgprotoporphyrin IX methyltransferase; CRD1, Mg-protoporphyrin IX monomethylester cyclase;
POR, NADPH:protochlorophyllide oxidoreductase; DVR, divinyl-protochlorophyllide
reductase; CHLG, chlorophyll synthase; CAO, chlorophyllide a oxygenase. Figure drawn
according to Mochizuki et al. (2010).
24
Introduction
The enzymes of the chlorophyll biosynthesis pathway from glutamyl-tRNA to
protoporphyrinogen are localized in chloroplast stroma (Joyard, et al., 2009).
Subsequent enzymes from protoporphyrinogen oxidase through to POR are associated
with both chloroplast envelope and thylakoid membranes. However, the localization of
Mg-chelatase is puzzling, since GUN4 and different subunits of Mg-chelatase have
been found in both soluble and membrane-containing fractions of purified chloroplast
(Guo, et al., 1998; Nakayama, et al., 1998; Larkin, et al., 2003; Joyard, et al., 2009)
CHLI is localized in the stroma and CHLD has been suggested to be localized
predominantly in stroma but to some extent in thylakoid membranes as well (Guo, et
al., 1998; Joyard, et al., 2009). CHLH is localized in both stroma and envelope
membranes (Nakayama, et al., 1998). Moreover, Mg-chelatase has been reported to
respond to Mg2+ concentration and to associate with envelope membranes when Mg2+
concentration is high but to dissociate from membranes when Mg2+ concentration
decreases (Guo, et al., 1998; Nakayama, et al., 1998). Of the enzymes catalyzing the
final steps of chlorophyll biosynthesis, chlorophyll synthase is localized in thylakoid
membranes (Joyard, et al., 2009) but chlorophyllide a oxygenase (CAO) has been
suggested to be present in envelope membranes (Eggink, et al., 2004).
Chlorophyll molecules that are not properly assembled to protein complexes are able to
direct the light energy to unappropriate targets, such as molecular oxygen, which
results in the formation of ROS that can harm the cell constituents. Since chlorophyll is
synthesized in particularly large amounts during chloroplast biogenesis it is clear that
chlorophyll biosynthesis is tightly regulated. The regulation is achieved by strong
transcriptional control of specific genes together with post-translational regulation in
chloroplasts which allow rapid changes in the flow rate of the pathway. HEMA1,
CHLH, CRD1 and CAO have been demonstrated to comprise a group of genes that
strongly respond to light and circadian signals (Matsumoto, et al., 2004). The ratelimiting step in the pathway is the activity of glutamyl-tRNA reductase and the
synthesis of ALA, which is feedback-inhibited by the accumulation of the end products
of both heme branch and chlorophyll branch. Feedback-inhibition of HEMA1 in
response to accumulation of protochlorophyllide in the chlorophyll branch is achieved
via activation of FLU that suppresses the activity of HEMA1 (Meskauskiene, et al.,
2001; Goslings, et al., 2004). CHLI, one of the three subunits of Mg-chelatase
(Ikegami, et al., 2007), has been identified as a target for thioredoxin-dependent
regulation. Also the activity of CAO is modified post-translationally as it is
destabilized in response to accumulation of chlorophyll b (Yamasato, et al., 2005).
1.5. Regulation of chloroplast biogenesis and acclimation to environmental
cues
The chloroplast differentiation process is modulated by environmental cues, of which
the most important is light, and it depends on coordinated action of nuclear and
organellar gene expression. To sense light, plants use a range of different
Introduction
25
photoreceptors that cover a broad spectrum of light signals. Sensitivity to red and farred light is mediated by phytochromes, the family of which contains five proteins
(phyA-phyE) in Arabidopsis. Two of these, phyA and phyB, are particularly important
during de-etiolation, the conversion of an etioplast to chloroplast (Reed, et al., 1994;
Mazzella, et al., 2001). Blue light is sensed by cryptochromes (cry1 and cry2 in
Arabidopsis; Cashmore, et al., 1999), of which cry 1 has a dominant role during deetiolation under blue light (Mazzella, et al., 2001).
1.5.1. Anterograde signaling
Since chloroplast biogenesis is tightly bound to the development of its ‘host’ cell and
largely dependent on the import of nuclear-encoded proteins, communication between
nucleus and chloroplast is required in order to adjust gene expression in both
compartments to ensure co-regulation of genes whose products function together.
Anterograde (nucleus-to-organelle) mechanisms coordinate gene expression in
organelles in response to endogenous and environmental signals perceived by the
nucleus. Nuclear-encoded proteins primarily control the gene expression in the
plastids, mostly by post-transcriptional mechanisms (Woodson and Chory, 2008), but
also chloroplast division and differentiation are dependent on nuclear-encoded
proteins. Chloroplasts of higher plants contain two RNA polymerases, nuclear-encoded
one (NEP) and plastid-encoded one (PEP) whose activity is, however, fine-tuned by
nuclear-encoded sigma factors (Maliga, 1998). The two RNA polymerases and
different nuclear-encoded sigma factors are essential during different stages of
Arabidopsis development (Kanamaru and Tanaka, 2004). It is assumed that NEP and
PEP act sequentially during the chloroplast development: NEP is active in proplastids
transcribing housekeeping genes, and PEP initiates the transcription of photosynthesisrelated genes and takes over the transcription of housekeeping genes in developing
chloroplasts (Maliga, 1998). Of the 6 Arabidopsis nuclear-encoded sigma factors, SIG2
is essential for chloroplast development as it has a role in transcription of glutamatespecific tRNA gene. Thus SIG2 has the potential for controlling the flux of glutamyltRNA for both protein synthesis and chlorophyll synthesis in chloroplasts. SIG5
instead responses to various stress conditions and contributes to the repair of the
damaged PSII reaction center proteins (Kanamaru and Tanaka, 2004).
1.5.2. Chloroplast-to-nucleus retrograde signaling
In order to maintain mature chloroplasts optimally functional, chloroplast operations
need to be adjusted according to the environmental cues. Changes in the light
conditions result in changes in the photosynthetic flux, and damage of chloroplasts by
high light or pathogens induces repairing mechanisms. Coordinated regulation of gene
expression in the nucleus and plastids is crucial for the acclimation of plants as the
26
Introduction
intracellular communication between organelles establishes the proper balance of gene
expression products in a changing environment.
Retrograde (organelle-to-nucleus) signalling plays a key role in optimizing plastid
functions. Retrograde mechanisms transmit signals that have originated in the
organelle to regulate gene expression in nucleus that can then modify anterograde
signals. Thus retrograde signals provide information of the metabolic and
developmental stage of organelles for the nucleus, and it is now obvious that these
signals are produced by several different processes in plastids, including
photosynthesis, pigment biosynthesis and metabolism (Beck, 2005; Nott, et al., 2006;
Piippo, et al., 2006). Plastidial signals can be classified into several groups depending
on where they originate from: (i) redox processes in photosynthesis, (ii) pigment
biosynthesis, (iii) metabolite pool changes, and (iv) generation of ROS.
The redox state of photosynthetic electron transport chain and the levels of ROS that
are continuously formed as byproducts of photosynthesis depict the chloroplast’s
photosynthetic performance which needs to be adjusted according to environmental
changes. Relevant redox-active components that can be a source of signal are the PQ
pool and the PSI acceptor site molecules (NADPH, thioredoxin and glutaredoxin)
(Baier and Dietz, 2005). However, it seems unlikely that either ROS or redox
compounds could themselves act as signalling molecules that traverse to cytosol.
The chlorophyll biosynthesis intermediate, Mg-protoporphyrin IX, has been considered
to act directly as a signalling molecule (Strand, et al., 2003) and to travel to cytosol
(Ankele, et al., 2007), but recent precise and reproducible experiments have questioned
the hypothesis (Mochizuki, et al., 2008; Moulin, et al., 2008). Nevertheless, studies
with a unicellular red alga Cyanidioschyzon merolae have shown that nuclear DNA
replication, which precedes cell division, is regulated by tetrapyrrole signals, namely
protoporphyrin IX and and Mg-protoporphyrin IX (Kobayashi, et al., 2009).
Furthermore, another composer of chlorophyll biosynthesis, H-subunit of Mgchelatase, might still mediate plastid signalling as was suggested before by
(Mochizuki, et al., 2001). Nonetheless, protein export from chloroplasts has not been
demonstrated and the protein import apparatus is known to act unidirectionally (Kleine,
et al., 2009).
Metabolites, especially carbohydrates, are strong candidates for signalling molecules as
photosynthesis is tightly integrated with cellular metabolism. It is well known that
increased levels of photosynthetic end products, glucose and sucrose, repress the
expression of photosynthetic genes (reviewed by Rolland, et al., 2006). Moreover,
small amounts of hexokinase, a cytosolic enzyme important for sensing and responding
to intracellular glucose signals, are found in the nucleus (Cho, et al., 2006) thus
implying a direct metabolic connection between photosynthesis and nuclear gene
expression.
Introduction
27
1.5.3. ROS in plant signaling
ROS are toxic compounds but they are also involved in signaling networks in plants.
Singlet oxygen, superoxide and hydrogen peroxide produced in chloroplasts are
implicated to participate in chloroplast-to-nucleus retrograde signaling. The connection
between H2O2 and signaling networks has been well-documented for many biotic and
abiotic stress responses (Larkindale and Knight, 2002; Apel and Hirt, 2004; Mateo, et
al., 2006). In fact, H2O2 is the ROS that has been recognized to induce the largest
changes in the levels of gene expression in plants, and this is probably due to its
relative stability (Dat, et al., 2000; Bechtold, et al., 2008; Fahnenstich, et al., 2008;
Foyer and Noctor, 2009). Yet it is not clear whether H2O2 is actually the signal itself,
or whether H2O2 oxidizes other molecules to generate an intracellular signal (Desikan,
et al., 2004). In microbes, proteins with H2O2 sensing capabilities have been identified
(Lee and Helmann, 2006; Toledano, et al., 2004; Manchado, et al., 2000; Zheng, et al.,
1998). These sensors share a mechanism in which H2O2 reacts with a unique cysteine
residue, coupling H2O2 metabolism and thiol redox signaling. To date, no H2O2
receptor has been definitively identified in plants (Foyer and Noctor, 2009). However,
signal molecules are usually present in cells in very low amounts, and plant cells seem
to tolerate higher concentrations of H2O2 than animal cells. The endogenous
concentration of H2O2 in plant cells is reported to range from nanomoles to several
hundred micromoles of H2O2 per gram fresh mass (Willekens, et al., 1997; Karpinski,
et al., 1999; Veljovic-Jovanovic, et al., 2002), while H2O2 is toxic for most animal cells
at levels of about 10–100 μM (Slesak, et al., 2007). It has been suggested that plants
tolerate high H2O2 levels due to the fact that plant antioxidant response systems are
designed more for the control of the cellular redox state than for complete elimination
of H2O2 (Slesak, et al., 2007). Superoxide and singlet oxygen are highly reactive
compounds and therefore possibly able to bind and modify the activity of some kinases
and phosphatases (Reinbothe, et al., 2010). Superoxide and singlet oxygen
accumulating in chloroplasts during photosynthesis have been proposed to trigger
refined signaling cascades involving plant hormones that modify plant growth
(Reinbothe, et al., 2010; Pilon, et al., 2011). However, both superoxide and singlet
oxygen produced in chloroplasts have only limited ability to cross membranes thus
being unable to affect signaling pathways outside of the chloroplast (Apel and Hirt,
2004).
The limitation of H2O2 and other ROS as signaling compounds is the fact that they lack
specificity; they are simple molecules, unable to store or transmit complex information,
and most of them are very reactive and thus short-lived. It has been proposed that
rather than ROS themselves, peptides derived from degradation of ROS-damaged
proteins could be more specific and selective secondary messengers to the nucleus
(Møller and Sweetlove, 2010). Nonetheless, due to its dual role as a damage-inducing
and a signal-inducing compound, the levels of H2O2 in the cells must be carefully
balanced.
28
Aims of the study
2. Aims of the study
Plants contain a particularly large number of thioredoxin isoforms in different cellular
compartments, including the cytosol, nucleus, mitochondria and chloroplasts.
Chloroplasts are different from other cell organelles since in addition to numerous
thioredoxin isoforms located to chloroplasts, they also contain two different
thioredoxin reductase systems. Chloroplasts are also unique because of the high
production of ROS in light-dependent reactions, and thioredoxins are closely linked
with the antioxidant systems scavenging hydrogen peroxide. In my thesis work, I
aimed i) to identify chloroplast processes regulated by the NADPH-dependent
thioredoxin system involving NTRC, the thioredoxin reductase that was recently
discovered when the experimental work for my doctoral thesis was about to start; and
ii) to reveal the impact of NTRC and other chloroplast antioxidant enzymes on the
acclimation of plants to various light regimes.
Methodological aspects
29
3. Methodological aspects
3.1. Plant material and growth conditions
Arabidopsis thaliana ecotype Columbia plants were used in all experiments. T-DNA
insertion mutants (Alonso et al. 2003) ntrc (At2g41680; SALK_096776 and
SALK_114293), sapx (At4g08390; SALK_083737) and tapx (At1g77490;
SALK_027804) were obtained from the Salk Institute (http://signal.salk.edu/). The
homozygosity of the mutants was assessed by PCR with two gene-specific primers and
one primer specific to T-DNA insertion. A tapx sapx double mutant was created by
crossing tapx and sapx single mutants, and was identified from the F2 generation by
PCR analysis by using the same set of primers which were used to identify the single
mutants.
Plants were grown at 23 °C under moderate white light (growth light; 100-150 µmol
photons m-2 s-1) in 8-h light/16-h dark periods (short-day-plants) (Papers I-IV), 16-h
light/8-h dark periods (long-day-plants) (Papers I-III), continuous light (Papers I-II) or
several distinct light-dark periods (Papers II-III). In Paper IV, plants were treated with
high light intensities (300-1300 µmol photons m-2 s-1) or low temperature (10 °C) for 2
h to 2 weeks. In Paper I, plants were also germinated under different spectral qualities
of light: red (30 µmol photons m-2 s-1), blue (3 and 30 µmol photons m-2 s-1) and far-red
light. Mature leaves or rosettes of 3-4 weeks old Arabidopsis plants were used as
material in most of the experiments in Papers I-IV. 5-day-old and 10-day-old seedlings
were also used in Paper I, and 7-day old seedlings in Paper IV.
3.2. Stress treatments
For short-term stress treatments, 4-week-old plants grown under growth light and an 8h photoperiod were shifted to higher light intensities, described in more detail in Paper
IV, for 2 to 6 hours. For long term stress treatments, plants were first grown under
growth light for 2 weeks, and thereafter shifted to higher light intensities or low
temperature, described in more detail in Paper IV, for 2 weeks.
The sensitivity of mature rosettes to methyl viologen -induced photo-oxidative stress
was explored by spraying the plants with 50 µM methyl viologen at the end of the dark
period. In the subsequent morning, plants were illuminated under growth light or high
light as described in the Paper IV. The extent of cell death and membrane disruption
was measured as ion leakage of excised rosettes to ion-exchanged water with a
conductivity meter (Radiometer).
30
Methodological aspects
The sensitivity of germination to photo-oxidative stress was explored by germinating
seeds on filter paper on Petri dishes containing half-strength Murashige and Skoog
medium supplemented with 1,5 µM methyl viologen and following greening under
growth light.
3.3. Analysis of pigments and proteins
Chlorophyll content of leaf disks was determined in DMF (dimethylformamide)
according to Inskeep and Bloom (1985). Anthocyanin content of leaf disks was
measured according to Neff and Chory (1998) with small modifications described in
Paper I. Isolation of thylakoid membranes and total root and leaf extracts was
performed as described in Paper I. Chlorophyll content of isolated thylakoids was
measured in HEPES-buffered acetone according to Porra et al. (1989), and the protein
contents of the total and soluble extracts were determined with the Bio-Rad Protein
Assay Kit. Proteins were separated with 12% (w/v) SDS-PAGE and detected using
specific antibodies described in the respective papers.
3.4. Microarray analysis
For microarray analysis, total RNA was isolated with Trizol reagent and labeled by the
aminoallyl method with Cy3 or Cy5 fluorescent dyes. RNA isolation, cDNA synthesis,
labeling, hybridization and the data analysis were performed as described in detail in
Papers I and IV.
3.5. Biophysical methods
The photoinhibition state of PSII was monitored as a ratio of variable to maximal
chlorophyll fluorescence, Fv/Fmax (Fv is a difference between maximal, Fmax, and
initial, Fo, fluorescence), measured from intact leaves with a Hansatech PEA
fluorometer after a 30 min dark incubation. Gas exchange of intact Arabidopsis plants
was measured with CIRAS-1 combined IR gas analysis system (PP Systems) equipped
with an Arabidopsis pot chamber (PP Systems). The response of net photosynthesis
(An) to the reference CO2 was measured under PPDF (photosynthetically active photon
flux density) that was saturating for net photosynthesis (500 μmol m−2 s−1 at 20 °C).
The parameters for maximal carboxylation rate of Rubisco (Vcmax, μmol of CO2 m−2
s−1), maximal electron transport rate (Jmax, μmol m−2 s−1) and rate of mitochondrial
respiration in light (Rd, μmol m−2 s−1) were obtained by modelling the response of net
CO2 assimilation to increasing extracellular CO2 concentration according to Farquhar
et al. (1980).
Methodological aspects
31
3.6. Microscopy
Confocal microscope images in Papers I and IV were obtained with an inverted
confocal laser-scanning microscope (Zeiss LSM510 META) using a 20x/0.50 water
objective. Chlorophyll fluorescence was excited at 488 nm with an argon diode laser,
and detected with a 650-710 nm passing emission filter. Maximal projections of
sequential confocal images were created with the Zeiss LSM Image Browser software.
The samples for light microscopy and electron microscopy were prepared according to
Pätsikkä et al. (2002). Electron microscope images were obtained with a transmission
electron microscope (JEOL JEM-1200EX).
3.7. In vivo-detection of H2O2 and superoxide
Accumulation of H2O2 and superoxide in the leaves was detected using DAB
(diaminobenzidine; Sigma–Aldrich) and NBT (nitroblue tetrazolium, Sigma-Aldrich),
respectively, as substrates. Rosettes were excised at the end of the light period, and
incubated on Petri dishes containing a solution of DAB or NBT overnight in darkness.
In the subsequent morning, the dishes were transferred to growth light for 1 h and
thereafter the rosettes were incubated in ethanol until chlorophyll was bleached. Finally
the rosettes were photographed.
3.8. Analyses of amino acids, hormones and sugars
Plant material for amino acid analysis was treated and extracted as described in Paper
I. Amino acid content of the extracts were further extracted and derivatized with the
Ez:faast liquid chromatography-mass spectrometry kit (Phenomenex) using the
procedure described by Husek (1998) and analyzed as propyl chloroformates with
HPLC-ESI/MS. Plant hormones were analyzed in Paper I by gas chromatography-mass
spectrometry analysis using a modified vapour-phase extraction method described by
Schmelz et al. (2003). Soluble sugars were extracted from the plant material as
described in Paper III. Sucrose, D-fructose and D-glucose were measured
spectrophotometrically with Sucrose/Fructose/D-Glucose assay kit (Megazyme). For
measurement of starch content, the remaining leaf material was dried and ground in
liquid nitrogen. Starch was solubilised and the total starch content was measured
spectrophotometrically with Total Starch assay kit (Megazyme).
32
Methodological aspects
3.9. Yeast two-hybrid analysis
The yeast strain CY306 (Vignols, et al., 2005) that carries deletions of endogenous
genes encoding cytosolic TRXs was used throughout the yeast two-hybrid
experiments. Escherichia coli DH5α –strain was used in molecular cloning. For the
yeast two-hybrid prey constructs, Riken RAFL (Riken, BRC, Japan) cDNA clones
were used as templates to generate PCR products from open reading frames of
Arabidopsis genes and to clone them cloned in frame with the activator or binding
Gal4 domains in pGAD.T7 and pGBK.T7 vectors (Clontech). For the yeast two-hybrid
bait constructs, full length open reading frame of NTRC gene or NTRC gene construct
including mutated cysteines (C217S, C220S, C457S) was used as a template in PCR to
generate truncated NTR and TRX domains. Double transformants in CY306 strain
were selected as the cells were grown on YNB -agar Petri dishes in the presence of
histidine, uracile, lysine, adenine and methionine but in the absence of leucine and
tryptophan. Cells bearing interacting proteins were further selected on the medium
lacking tryptophan, leucine and histidine.
Overview of the results
33
4. Overview of the results
4.1. Daily light/dark rhythm modulates the photosynthesis and growth of
Arabidopsis
Characterization of the ntrc lines lacking chloroplast NADPH-dependent thioredoxin
reductase demonstrated that the mutant phenotype strongly depended on the
photoperiod, to which plants were acclimated during the growth (Paper I). Therfore I
studied in detail how the daily light/dark rhythm modulates the growth and metabolism
of Arabidopsis in general (Papers I-III). The results in this thesis demonstrate that like
acclimation to various light intensities and CO2 concentrations, daily photoperiods
modulate the photosynthetic structures of wild type Arabidopsis leaves and induce
changes in the vegetative growth of plants.
4.1.1. Impact of photoperiod on growth and photosynthesis in wild type
Arabidopsis
The growth rate of Arabidopsis is determined by the efficiency to convert the
assimilated carbon to biomass (Zeeman et al. 1998, Gibon et al. 2004). Like low light
intensity and low CO2 concentration, short daily photoperiods decreased the
photosynthetic carbon assimilation and resulted in deceleration in the vegetative
growth of plants (Paper I). In Papers II and III, the effect of the photoperiod on the
growth of Arabidopsis was further studied by growing plants in continuous light and
under various light-dark regimes, some of which contained an interrupted dark period
or was deviated from the natural 24-h rhythm. Short photoperiods in combination with
long nights reduced the growth of Arabidopsis rosettes, measured as a reduction in
biomass production (Paper III). Light-dark cycles deviating from the natural 24-hperiod did not diminish the growth when the dark period was short. Furthermore, long
photoperiods were not able to compensate for the disadvantages caused by long dark
periods (10 h light/10 h dark, 12 h light/12 h dark vs. 16 h light/16 h dark). Thereby the
reduction in growth is not determined by the total duration of the light available for
photosynthesis during the growth, but instead by the changes in metabolism induced by
the light-dark rhythm itself. Accordingly, slow growth rates observed under short
photoperiods correlate well with the efficiency of starch metabolism; especially with
the lower starch degradation rates under short photoperiods (see 4.2.2).
In this thesis work it is further demonstrated that similar to light intensity (reviewed by
Kim et al., 2005) also the length of the daily light/dark periods modulates the structure
of the leaf and the development of mesophyll cells and chloroplasts in Arabidopsis
34
Overview of the results
(Papers I and II). Also adjustments to the photosynthetic apparatus were observed that
are comparable to changes induced by light intensity (Papers I-II). Arabidopsis wild
type plants grown under short photoperiods with long nights resemble low-light-grown
plants with poorly developed, roundish palisade mesophyll cells and thin leaves. On
the contrary, plants grown under long photoperiods or continuous light resemble highlight-acclimated plants with thick leaves and well-developed, tightly packed and
vertically elongated palisade mesophyll cells (Paper II; Yano and Terashima, 2001). In
addition to adjustments in the anatomy of the palisade mesophyll, an increase of 40%
in the stomatal index, which refers to the number of stomata per leaf area, was
observed in plants grown under long photoperiods. As the gas exchange and
transpiration through stomata are crucial determinants of the photosynthetic
performance of plants, also the net CO2 assimilation per rosette was 20 to 40 % higher
in long-day-grown leaves than in short-day-grown leaves at both ambient CO2
concentration and in saturating light intensity (Paper I). Plants grown under long days
have also higher chlorophyll content per leaf area due to thicker leaves and a higher
chlorophyll a/b ratio when compared to plants grown under short days (Paper I). High
chlorophyll a/b ratio of long-day-grown leaves in comparison to short-day-grown
leaves implies that the photoperiod has an influence on the composition of the lightharvesting complexes in the thylakoid membranes. Indeed, the grana stacks were
smaller in chloroplasts of plants grown under long days (Paper II) and accordingly, the
relative amount of the trimeric chlorophyll a/b-binding proteins of the PSII antenna in
thylakoid membranes was smaller in long-day-grown plants when compared to shortday-grown plants. However, no significant difference was observed in the relative
proportion of the representative subunits of PSII, PSI and Cyt b6f complexes in plants
grown under short or long photoperiods (Paper I).
4.1.2. Impact of photoperiod on the redox metabolism of mesophyll cells
As shown in Papers I-II, different light-dark rhythms induce adjustments in the
photosynthetic structures in Arabidopsis leaves. The underlying mechanism for such
modifications remains to be resolved, but the length of the photoperiod also induced
notable changes in ROS metabolism (Paper III). Short photoperiods increased the
production of ROS in Arabidopsis leaves (Paper III), as especially H2O2 accumulated
in the leaves during the light periods. Also thylakoids isolated from short-day-grown
plants produced more ROS than thylakoids isolated from long-day-grown plants (Paper
III). However, the analysis of the steady-state levels of ROS scavenging enzymes
revealed that no significant compensatory changes occur in the accumulation of
chloroplast antioxidative enzymes. Only the amount of chloroplast PrxIIE was slightly
higher in plants grown under short photoperiods compared to those under long
photoperiods. Apparently the antioxidative systems in chloroplasts are not able to
intensify to counteract the increased production of ROS under short photoperiods. On
the other hand, the production of ROS may also be of physiological expedient,
standing for the regulation of metabolism during acclimation of plants to different
light-dark rhythms. If the changes in the redox homeostasis are crucial to induce
Overview of the results
35
acclimation of plant to various light regimes, thioredoxins that are abundant in
chloroplasts can be considered as significant regulatory components in the acclimation
processes.
4.1.3. Knockout of NTRC impairs the acclimation of Arabidopsis to short
photoperiods
Plants lacking chloroplast NADPH-dependent thioredoxin reductase showed more
distinct photoperiod-dependent phenotypes than wild type Arabidopsis. Indeed, the
growth rate, biomass production and chlorophyll accumulation varied strongly in ntrc
plants under different daily light-dark-cycles (Papers I-III). Cotyledons of ntrc plants
were visually indistinguishable from those of wild type plants, but upon the emergence
of the first true leaves the mutant phenotype became evident especially under short
photoperiods. During the first month of growth under 8-hour photoperiod, ntrc plants
formed small rosettes with pale green leaves and had a low chlorophyll content. Upon
ageing, the ntrc leaves started to green and eventually the rosettes gained the size
similar to mature wild type plants. Moreover, the flowering time of ntrc plants was
significantly delayed under short photoperiods (Paper I). In general, the chlorophyll
accumulation in wild type plants was significantly lower upon the short 8-hour
photoperiod, suggesting that the length of the photoperiod is a critical factor regulating
chlorophyll accumulation. In plants deficient in NTRC, such reduction of chlorophyll
accumulation under short photoperiods, less than nine hours, was even more
pronounced than in wild type plants (Papers I-II). Low chlorophyll content was
associated with reduced number of chloroplast per ntrc mesophyll cell (Papers I-II).
Also the biomass production was more prominently reduced in ntrc plants grown under
short photoperiod with long nights than in wild type plants (Paper III). The growth
under longer photoperiods, 16 h light/8h dark or continuous light, enhanced the
accumulation of both chlorophyll and biomass as well as the growth rate in ntrc plants.
Also, the transition to flowering took place approximately at the same time in ntrc and
in wild type plants (Paper I). Thus it appears that NTRC is essential for the proper
acclimation of Arabidopsis to short photoperiods combined with long nights.
4.1.4. Photosynthesis of knockout ntrc lines
Low chloroplast number per cell and the occurrence of abnormal chloroplasts in ntrc
plants, especially under short photoperiod, was accompanied with low CO2 fixation
capacity and high CO2 compensation point of photosynthesis (Paper I). The ntrc plants
grown under short photoperiod suffered also from enhanced photoinhibition of PSII,
measured as a decrease in Fv/Fm in growth light intensity. Consistent with the high CO2
compensation point, ntrc lines had doubled respiration rate when compared to Col-0
leaves (Paper I). The transcript profiling of short-day-grown ntrc plants also showed
induction of several photorespiratory genes (Paper I) that suggests an elevated
36
Overview of the results
utilization of light energy in photorespiration. Under long photoperiod, the differences
in the net CO2 assimilation, CO2 compensation point and photoinhibition of PSII
between ntrc and wild type plants were less distinct. However, the rate parameters of
photosynthesis and the pattern of photosynthetic thylakoid membrane protein
complexes in ntrc grown under both short and long photoperiods was comparable to
that of wild type plants (Paper I) implicating that the basic structure and function of
light and carbon fixation reactions were not affected by the knockout of NTRC enzyme
in Arabidopsis.
4.2. Identification of chloroplast processes controlled by NADPHdependent thioredoxin system
The results in this thesis demonstrate that the thioredoxin system has higher impact on
the development, metabolism and acclimation of chloroplast than was previously
assumed in the light of studies with ferredoxin-thioredoxin system. The two
thioredoxin systems in plant chloroplasts, the ferredoxin-dependent system and the
NADPH-dependent system, have distinct roles in redox-regulation of chloroplast
functions. By comparing the growth, responses to environment, biochemical properties
and gene expression in wild type plants and plants lacking chloroplast NADPHdependent thioredoxin reductase, I conclude that NTRC regulates chloroplast
biogenesis and carbon metabolism subsequent to Calvin-Benson cycle such as starch
metabolism and shikimate pathway, as well as is involved in defence reactions against
oxidative stress. On the contrary, ferredoxin-thioredoxin system seems to be the key
regulator of primary photosynthetic reactions such as Calvin-Benson cycle and redoxregulated enzymes therein.
4.2.1. Impact of NTRC on chloroplast biogenesis
The characterization of the ntrc knockout lines demonstrated that NTRC is involved in
regulation of chloroplast biogenesis. The chloroplast number in palisade mesophyll
cells was reduced in mature ntrc leaves, especially in plants acclimated to short
photoperiods (Papers I and II). The chloroplasts were also heterogeneous in size.
Transmission electron micrographs demonstrated developmental disorders in the
ultrastructure of chloroplasts in ntrc cells (Paper II). Some of the ntrc chloroplasts were
malformed and irregularly elongated in shape, exhibiting protrusions devoid of
thylakoid membranes. Also the distribution of the thylakoid membranes inside
chloroplasts was varying as several types of chloroplasts were observed in ntrc cells:
regular chloroplasts containing grana and stroma thylakoids identical to wild type
plants, chloroplasts with central areas well occupied with grana and stroma thylakoids
but one or both ends deficient in internal membranes and chloroplasts with only few
thylakoid membranes. Importantly, a population of chloroplasts with dissimilar
structure/function exists in a single ntrc mesophyll cell (Paper II). The different types
Overview of the results
37
of chloroplasts appeared often in a series in a single cell in which unimpaired plastids
resided next to chloroplasts with unbalanced distribution of thylakoids, and the plastids
with significant reduction in thylakoid membranes being situated at the end of the
chain. The largest number of anomalous chloroplasts was detected in mature ntrc
leaves grown under short photoperiods, but they were also observed already in the 5day-old cotyledons, in the young developing leaves, and in the mature leaves of plants
grown under long photoperiods or under continuous light (Paper II). The increasing
appearance of abnormal chloroplasts in maturating ntrc leaves implies that NTRC is
essential for the proper biogenesis of chloroplasts, possibly via regulation of
chloroplast division, chlorophyll biosynthesis and thylakoid formation.
FtsZ proteins form a core component of the inner division machinery in chloroplast
(Yang, et al., 2008). They are tubulin-like GTPases that form the ring structure on the
stromal surface of the envelope (Miyagishima, et al., 2006; Schmitz, et al., 2009). FtsZ
ring functions as a scaffold structure, on which the other division components assemble
(Maple, et al., 2005; Miyagishima, et al., 2006; Glynn, et al., 2008). Unlike
cyanobacteria that have only one FtsZ protein, plants have two FtsZ proteins, called
FtsZ1 and FtsZ2 (Osteryoung, et al., 1998). Both proteins localize at the division site,
and loss of either protein impairs chloroplast division (Schmitz, et al., 2009).
Interestingly, FtsZ has been identified by thioredoxin proteomics (Balmer, et al., 2003),
suggesting that thioredoxins may control the formation of chloroplast division
machinery. To get further insights into this assumption the interaction of NTR and
TRX domains of NTRC with FtsZ1 and FtsZ2 was tested by yeast two-hybrid
approach. In the test, an interaction between FtsZ1 and the Trx domain of NTRC was
observed (Paper II).
Besides the reduced number of chloroplasts in ntrc cells and the developmental
disorders in the ultrastructure of chloroplasts, also chlorophyll biosynthesis was
impaired in ntrc plants. Short light periods decreased significantly the accumulation of
chlorophyll in ntrc leaves in comparison to wild type plants (Papers I-II). In Paper I,
comparative transcript profiling of ntrc and wild type plants revealed alterations in the
expression of specific genes relating to chlorophyll biosynthesis. Among the most upregulated genes in short-day-grown ntrc leaves were two genes encoding the key
enzymes of the chlorophyll biosynthesis pathway, glutamyl-tRNA reducatse (HEMA1)
and the H-subunit of Mg-chelatase (GUN5), suggesting an unbalanced biosynthesis of
chlorophyll in ntrc leaves. A yeast two-hybrid approach was used to test if NTR or
TRX domains of NTRC interacts with chlorophyll biosynthesis enzymes HEMA1, Mgprotoporphyrin IX methyltransferase and subunits of Mg-chelatase, GUN4 and Isubunit. The analysis revealed no interaction between the I-subunit of Mg-chelatase
and the thioredoxin domain of NTRC (Paper II). HEMA1, Mg-protoporphyrin IX
methyltransferase and GUN4 showed only weak interaction with the thioredoxin
domain of NTRC (Paper II; Jouni Toivola, unpublished results).
The comparative transcript profiling revealed also changes in the expression of genes
involved in the thylakoid formation. The ntrc plants showed lowered accumulation of
transcripts for THYLAKOID MEMBRANE ORGANIZATION-LIKE (FZL) and
38
Overview of the results
THYLAKOID FORMATION 1 (THF1) as well as down-regulation of the expression of
LHCB1, LHCB2 and LHCB3 that encode for protein components of the lightharvesting complex of PSII. The reduced accumulation of LHCB, FZL and THF1
transcripts may be a consequence of the impaired chlorophyll biosynthesis, since the
proper formation of thylakoid membranes requires the simultaneous assembly of
membrane lipids, protein components and pigments to attain functional, threedimensional structures.
4.2.2. Starch and sucrose metabolism in knockout ntrc leaves
The acclimation of Arabidopsis to different photoperiods was accompanied by altered
sugar metabolism in vegetative leaves (Paper III). Wild type Arabidopsis plants grown
under short days had a higher rate of starch synthesis during the first hour of light
period and a lower rate of starch degradation in night than plants grown under long
days. Sucrose accumulated with the same rate under both short and long photoperiods,
the amounts left after dark period being lower in short-day-grown plants. The content
of hexoses, glucose and fructose increased in wild type plants during the first hours of
light period, but stayed rather stable or decreased towards the end of the day. The
content of glucose was similar in both short-day- and long-day-grown wild type plants
whereas the amount of fructose was lower in plants grown under short days.
The ntrc plants grown under both short and long photoperiods produced less sucrose,
glucose and fructose when compared to the wild type plants (Paper III). Under both
photoperiods, ntrc plants also accumulated less starch when compared to wild type
plants (Paper III). Partly it is due to decreased photosynthetic productivity caused by
the low chloroplast number per cell and the occurrence of abnormal chloroplasts,
especially in ntrc plants acclimated to short photoperiods. However, the reduced
apparent rates of starch accumulation was also detected in ntrc plants acclimated to
long photoperiods (Paper III), in which no reduction in chloroplast number was
observed (Paper II). Furthermore, the amount of starch in wild type plants in short
photoperiod began to rise directly after the onset of the light period and the
accumulation of starch was accelerated towards the end of the light period both under
short-day- and long-day-grown wild type Arabidopsis leaves. Under both conditions,
ntrc plants were not capable of accelerating starch biosynthesis in the end of the light
period, suggesting a requirement of NTRC in a redox control of enzymes in starch
synthesis. Also the degradation of starch during the first hour of subsequent dark
period was slightly slower in ntrc than in wild type plants under both photoperiods.
Nevertheless, no significant differences between ntrc and Col-0 were recorded in the
amount of starch left in the leaves after the dark period.
An ADP-glucose pyrophosphorylase (AGPase) and a dual specificity protein
phosphatase (DSP4) play important roles in the synthesis and degradation pathways of
starch, respectively (Neuhaus and Stitt, 1990; Sokolov, et al., 2006). Both enzymes are
also shown to be activated by thioredoxins (Ballicora, et al., 2000; Sokolov, et al.,
Overview of the results
39
2006). Furthermore, AGPase has been suggested to be regulated by NTRC (Michalska,
et al., 2009). Thereby the yeast two-hybrid approach was used to test if NTR or TRX
domains of NTRC interact with AGPase and DSP4. The analysis revealed an
interaction between the thioredoxin domain of NTRC and the small subunit of AGPase
(Paper III), but no specific interaction with DSP4 polypeptide and TRX of NTRC was
detected.
4.2.3. NTRC and chloroplast ROS metabolism
Earlier reports have shown that NTRC is able to reduce chloroplast 2-Cys Prxs
oxidized in the elimination reactions of H2O2 (Perez-Ruiz, et al., 2006; Kirchsteiger, et
al., 2009; Perez-Ruiz and Cejudo, 2009). In yeast two-hybrid screen (Paper III) a
strong interaction between NTRC and 2-Cys Prx was observed. The analysis of the
steady-state levels of ROS scavenging enzymes (Paper III) revealed that under normal
growth conditions, ntrc plants show distinctive modulations only in the levels of APXdependent and 2-Cys-Prx-dependent detoxification systems of H2O2. The protein level
of 2-Cys Prxs was significantly reduced and the levels of SOD and sAPX increased in
ntrc leaves grown both under short or long photoperiod when compared to wild type
plants (Paper III). 2-Cys Prxs are labile in ntrc plants, probably because of the
proteolytic degradation of the oxidized forms of 2-Cys Prx that cannot be reduced in
the absence of NTRC. Despite the shutdown of NTRC-dependent 2-Cys Prx system,
H2O2 or superoxide did not accumulate in significantly higher levels in leaves of
NTRC-deficient plants under normal growth conditions in comparison to wild type
Arabidopsis (Paper III). This can be explained by the compensation of 2-Cys Prx
system with ascorbate-dependent SOD and sAPX enzymes (Paper III) in the
detoxification of H2O2 in NTRC-deficient plants. Accordingly, knockout of both
chloroplast APXs induced an enhanced accumulation of 2-Cys Prxs in high-light
acclimated Arabidopsis leaves (Paper IV) indicating the tight interlinkage between the
ascorbate-dependent APX and NTRC-dependent 2-Cys Prx systems in the control of
H2O2 in chloroplast.
4.2.4. Knockout of NTRC interferes with the homeostasis of metabolites
produced in shikimate pathway
Aromatic amino acids are synthesized in chloroplasts via shikimic acid pathway and
they serve as precursors for the biosynthesis of auxin and flavonoids. Under short
photoperiods where the growth of ntrc plants is slowest, adjustments in the amount of
aromatic amino acids and metabolites derived from them were observed. i) The ntrc
seedlings grown under short photoperiod contained significantly less auxin than wild
type plants (Paper I). ii) ntrc plants were not capable of enhancing the accumulation of
anthocyanins in response to low temperature and senescence (Paper I). iii) The
accumulation of amino acids in the wild type and ntrc plants showed photoperiod-
40
Overview of the results
dependent modulations (Paper I). The amounts of glycine, alanine and threonine were
significantly lower in both wild type and ntrc plants grown under long photoperiods
than under short photoperiods. Aromatic amino acids tryptophan, phenylalanine and
tyrosine accumulated more in ntrc plants than in wild type plants, and the differences
were more pronounced under short photoperiods. Furthermore, under short
photoperiods, ntrc accumulated more arginine, asparagine and histidine while the
growth under long photoperiods diminished the differences.
In preliminary yeast two-hybrid analysis we tested if NTRC interacts with two
enzymes involved in shikimate pathway (Jouni Toivola, unpublished results). 3-deoxyd-arabino-heptulosonate-7-phosphate synthase (DAHP synthase) is the first enzyme in
shikimate pathway that is responsible for the formation of 3-dehydroquianate from
phosphoenolpyruvate and erythrose 4-phosphate, and tryptophan synthase β subunit
(TSB) catalyzes the conversion of indole to tryptophan which in turn can serve as a
precursor for auxin. Both enzymes have been identified as subjects for thiol-redoxregulation (Entus, et al., 2002; Balmer, et al., 2006; Kolbe, et al., 2006) and both
enzymes gave specific interaction with TRX-domain of NTRC (Figure 9, Jouni
Toivola, unpublished results).
Figure 9. Interaction of the TRX domain of NTRC with TSB2 and DAHP synthase in yeast
two-hybrid assay. Yeast CY306 cells co-expressing TRX domain of NTRC (native domain, or
domain carrying a mutation replacing the second cysteine of redox center CGPC by a serine) as
bait and target candidate (either TSB2, or DAHP synthase) proteins were grown into stationary
phase and adjusted to an OD600 of 0.5 and 0.05 before spotting onto plates containing histidine
(+HIS control panel) or without histidine (-HIS panel), respectively. Result indicates that both
TSB2 and DAHP synthase are able to interact with TRX domain of NTRC.
Overview of the results
41
4.3. Contribution of chloroplast ascorbate peroxidases to the ROS
metabolism in Arabidopsis
Arabidopsis plants contain five APX isoenzymes, two of which are located in
chloroplasts, thylakoid-bound APX (tAPX) and stromal APX (sAPX). In addition to
chloroplasts, sAPX is dual targeted to mitochondria as well. APXs have long been
considered as key enzymes in detoxification of H2O2, but their functional specificities
have been poorly understood. Characterization of single tapx and sapx mutant lines and
double tapx sapx mutant line of Arabidopsis (Paper IV) showed that sAPX is
particularly important for photoprotection during the early greening process in
germinating seedlings. In mature leaves, tAPX and sAPX are functionally redundant,
and crucial upon sudden onset of oxidative stress. The analysis of the steady-state
levels of other, possibly compensatory ROS scavenging enzymes in tapx and sapx
single mutant plants and tapx sapx double mutant plants revealed that under normal
growth conditions no distinct modulations in the levels of H2O2-detoxifying enzymes
were observable for tapx and sapx single mutant plants or tapx sapx double mutant
plants (Paper IV). Only high-light-stress-acclimated tapx sapx double mutant plants
showed increased level of 2-Cys Prxs, and low-temperature-acclimated wild type and
tapx plants showed upregulation of sAPX (Paper IV). As mentioned in the section
4.2.3., plants deficient in chloroplast APXs compensate for the lack of APXs by
upregulating 2-Cys Prxs, whereas plants deficient in NTRC upregulate SOD and sAPX
(Paper III). Thus the two H2O2-detoxifying systems in chloroplast, ascorbate-dependent
APX and NTRC-dependent 2-Cys Prxs are tightly linked up to prevent the detrimental
accumulation of ROS in chloroplast.
42
Discussion
5. Discussion
Plant thioredoxins were initially defined as regulators of the enzymes in Calvin-Benson
cycle and malate valve in chloroplasts (Schurmann and Jacquot, 2000), but the release
of Arabidopsis genome (The Arabidopsis Genome Initiative, 2000) and the
development of proteomic approaches (Verdoucq, et al., 1999; Motohashi, et al., 2001;
Balmer, et al., 2003; Maeda, et al., 2004; Marchand, et al., 2004; Hägglund, et al.,
2008) have allowed the identification of numerous novel thioredoxin targets in
chloroplasts as well as in cytosol, mitochondria and nucleus. Up to date, more than 400
thioredoxin target proteins have been identified in plants (Montrichard, et al., 2009)
and they are implicated in almost all aspects of plants’ life from housekeeping
metabolism to stress defense mechanisms. However, the implication of thioredoxin in
the functions of many thioredoxin target proteins identified by proteomic approaches
still remains to be resolved.
Plants contain two main types of thioredoxin reductases, ferredoxin-dependent
thioredoxin reductase (FTR) located in plastids and NADPH-dependent thioredoxin
reductases (NTRs) located in cytosol, mitochondria and chloroplasts (Laloi, et al.,
2001; Serrato, et al., 2004; Reichheld, et al., 2005). Originally, ferredoxin-thioredoxin
system was thought to be solely responsible for thiol-redox regulation in chloroplasts
but the discovery of chloroplastic NADPH-dependent thioredoxin reductase (NTRC;
Serrato, et al., 2004) has broadened the view of redox regulation of chloroplastic
proteins. Ferredoxin-thioredoxin system is dependent on ferredoxin reduced in light
during photosynthesis, while besides the light reactions, NADPH can be produced in
darkness via oxidative carbon metabolism. In this thesis, the role of NTRC in plant
development, acclimation and detoxification of ROS was studied. As shown in Papers
I, II and III, knockout of NTRC led to severe photoperiod- and age-dependent
developmental disorders that were especially pronounced under short photoperiods.
This is supported by an earlier report of Perez-Ruiz et al. (2006) that plants lacking
NTRC are hypersensitive to prolonged darkness, thus indicating that the two plastidial
thioredoxin reductases, FTR and NTRC, are functionally nonredundant.
5.1. NTRC in redox-regulation of chloroplast biogenesis
The reduced amount and altered ultrastructure of chloroplasts (Papers I-II) in plants
deficient in NTRC indicate that chloroplast differentiation is deteriorated in the
absence of NTRC. Since the formation of functional chloroplasts is vital for plant
survival, chloroplast differentiation requires tightly coordinated regulation of multiple
processes including chloroplast division, formation of thylakoid membranes,
chlorophyll biosynthesis and assembly of photosynthetic protein complexes. As
regards to thiol-redox regulation of these processes, three proteins in chlorophyll
biosynthesis and one protein in chloroplast division have been implicated as targets for
Discussion
43
thioredoxin-dependent regulation (Balmer, et al., 2003; Montrichard, et al., 2009),
supporting the hypothesis that NTRC has a role in controlling chloroplast biogenesis.
NADPH-thioredoxin system, rather than ferredoxin-thioredoxin system, is a
preferential candidate to regulate chloroplast biogenesis since metabolically produced
NADPH is available in differentiating chloroplasts prior the assembly of the light
reactions.
5.1.1. Regulation of chloroplast division by NTRC
Plants have two FtsZ proteins, FtsZ1 and FtsZ2 form heterodimers that polymerize to a
ring-like structure (Z-ring) at the chloroplast division site and both proteins are
essential for correct chloroplast division (reviewed by Miyagishima and Kabeya,
2010). FtsZ protein has been identified by thioredoxin proteomics (Balmer, et al.,
2003), suggesting that thioredoxins may be involved in regulating the chloroplast
division machinery. From the FtsZ proteins, an interaction between the thioredoxin
domain of NTRC and FtsZ1, but not with FtsZ2 protein, was observed in yeast twohybrid screen (Paper II). In a predicted amino acid sequence FtsZ1 protein contains a
concerved Cys that is missing in FtsZ2 protein (Paper II). If NTRC is involved in
regulating the formation of the FtsZ ring, the assembly of division machinery is
impaired in plants lacking NTRC resulting in lower accumulation of chloroplasts in
cells. Alternatively, the structural disorders in ntrc chloroplasts and the existence of
small plastid-like organelles in ntrc cells (Paper II) may refer to asymmetrical division
of chloroplasts. The correct position of the Z-ring in the middle of chloroplast is
mediated by a complex consisting of Min proteins and ARC3 protein (Maple and
Møller, 2007). FtsZ1 – but not FtsZ2 – has been shown to interact with Min/ARC3
complex (Maple and Møller, 2007). Min/ARC3 protein complex is localized at the
ends of a chloroplast, where the formation of the Z-ring is prevented by the binding of
the FtsZ1 to Min/ARC3 (Maple and Møller, 2007). The Min/ARC3 complex is absent
from the middle part of the chloroplast, thus allowing the Z-ring to assemble.
Accordingly, NTRC protein has been shown to be localized in clusters in the
chloroplast (Perez-Ruiz, et al., 2009). The role of NTRC in the regulation of
chloroplast division remains to be elucidated in the future but I hypothesize that a
disulphide bridge is formed between FtsZ1 and Min/ARC3 complex when they interact
with each other. The reduction of the disulphide bridge by a thioredoxin, presumably
NTRC, is needed to release the FtsZ1 protein from the complex and only a free FtsZ1
protein is capable to polymerize with FtsZ2 protein in the middle of a chloroplast. The
lack of a sufficient thiol reductant may disturb the division resulting in the
miscellaneous population of plastids in ntrc cell.
44
Discussion
5.1.2. Regulation of chlorophyll biosynthesis by NTRC
The characterization of ntrc mutant lines addressed to altered chlorophyll synthesis in
the absence of NTRC. The ntrc plants showed significant reduction in chlorophyll
content (Papers I-II; Perez-Ruiz, et al., 2006) and the transcript profiling of ntrc leaves
revealed an upregulation of specific genes coding for enzymes related to chlorophyll
biosynthesis (Paper I). Furthermore, NTRC gene expression has been shown to be coregulated with genes involved in chlorophyll biosynthesis (Stenbaek, et al., 2008). It
has been previously reported that the key enzyme in the chlorophyll branch of
tetrapyrrole biosynthesis, Mg-chelatase (reviewed by Stenbaek and Jensen, 2010) is
regulated by thioredoxin system (Balmer, et al., 2003; Ikegami, et al., 2007). The Mgchelatase consists of three subunits, CHLD, CHLH and CHLI, and is regulated by
GUN4 protein by an unknown mechanism (Larkin, et al., 2003). CHLI subunit
catalyses the hydrolysis of ATP that is stimulated by chloroplast thioredoxins
(Ikegami, et al., 2007). However, the thioredoxin-dependent regulation of Mgchelatase is still obscure, because thioredoxin treatment in vitro did not affect the
overall magnesium chelation activity of the enzyme (Ikegami, et al., 2007).
Accordingly, the yeast two-hybrid assay that failed to show any interaction between
NTRC and I-subunit of Mg-chelatase (Paper II) suggesting that Mg-chelatase is not a
target enzyme of NTRC. The weak interaction between GUN4 and NTRC in yeast
two-hybrid assay needs further elucidation before any conclusion can be drawn.
Stenbaek et al. (2008) showed that when plants were fed with ALA in darkness, the
intermediates of chlorophyll biosynthesis, Mg-protoporphyrin and Mg-protoporphyrin
monomethyl ester accumulated in higher amounts in ntrc than in wild type leaves,
suggesting that the reaction catalyzed by Mg-protoporphyrin IX methyltransferase
(CHLM) and Mg-protoporphyrin IX monomethylester cyclase (CRD1) (Figure 8) are
imbalanced in the absence of NTRC. A weak interaction between NTRC and CHLM
was detected in yeast two-hybrid assay, suggesting that NTRC may regulate the
activity of CHLM. Furthermore, slight increase in the activity of CHLM was observed
in vitro in the presence of NTRC (Anne Stenbaek, PhD thesis) that further supports the
thiol-redox regulation of CHLM enzyme. NTRC may also interfere indirectly with
chlorophyll biosynthesis enzymes by protecting enzymes from oxidation that
potentially occurs during chlorophyll synthesis (Stenbaek and Jensen, 2010). The latter
hypothesis is supported by in vitro experiments of (Stenbaek, et al., 2008) which
showed that in combination with 2-Cys Prx, NTRC was able to stimulate the activity of
CRD1, enzyme acting downstream of CHLM in the chlorophyll biosynthesis pathway
(Figure 8).
Discussion
45
5.1.3. Thylakoid formation in ntrc mutant lines
Transmission electron micrographs of chloroplasts demonstrated defective formation
of thylakoid membranes in ntrc mesophyll cells (Paper II). Microarray analysis on ntrc
(Paper I) also revealed a down-regulation of genes whose products act on formation of
thylakoid membranes, THYLAKOID MEMBRANE ORGANIZATION-LIKE (FZL) and
THYLAKOID FORMATION 1 (THF1) (Wang, et al., 2004; Gao, et al., 2006). During
the conversion of etioplasts to chloroplasts, thylakoid membranes initiate from
prolamellar bodies, tubular membrane aggregates arranged in a clustered manner inside
chloroplasts (recently reviewed by Solymosi and Schoefs, 2010). Localization of
NTRC protein in clusters in chloroplasts (Perez-Ruiz, et al., 2009) may alternatively be
linked with the redox-control of thylakoid formation by NTRC in chloroplasts.
Otherwise, the malformation of thylakoids observed in ntrc chloroplast may be a
secondary effect caused by the irregular plastid division and impaired chlorophyll
biosynthesis discussed in chapters 5.1.1. and 5.1.2.
5.2. Regulation of chloroplast metabolism by NTRC
5.2.1. Redox control of starch metabolism in chloroplasts
Transient formation of starch in light and degradation in darkness coordinates the
carbon assimilation and allocation to growth in Arabidopsis leaves (Zeeman, et al.,
2007; Paper III). The coordinated synthesis and remobilization of starch presumes the
regulatory signals from photosynthesis and respiration to the enzymes metabolizing
starch. The enzyme regulation is achieved via feed-back control by metabolites,
reversible protein phosphorylation and redox-regulation (Zeeman, et al., 2007; Kötting,
et al., 2010). ADP-glucose pyrophosphorylase (AGPase) is the key enzyme in starch
synthesis that controls the flux from carbon to starch. AGPase is a heterotetrameric
enzyme consisting of large and small subunits, which is redox-activated in light by the
reduction of a disulphide bridge between small subunits (Hendriks, et al., 2003).
AGPase is also allosterically controlled by the level of Pi and PGA, which act as an
inhibitor and as an activator, respectively (Zeeman, et al., 2007). Also the enzymes
involved in starch degradation, glucan, water dikinase (GWD), dual specificity protein
phosphatase (DSP4) and β-amylase 1 (BAM1) have been shown to be under redox
control (Mikkelsen, et al., 2005; Sokolov, et al., 2006; Sparla, et al., 2006). Prior to the
degradation by amylases, starch granules are reversibly phosphorylated by GWD and
DSP4 (Zeeman, et al., 2007). This reversible phosphorylation has been proposed to
disrupt the crystalline structure of amylopectin and mutant analyses have shown that
both enzymes are necessary to efficient remobilization of starch in Arabidopsis (Ritte,
et al., 2002; Yu, et al., 2001; Zeeman, et al., 2010). All these enzymes have been
46
Discussion
reported to be regulated by thioredoxins (Hendriks, et al., 2003; Mikkelsen, et al.,
2005; Sokolov, et al., 2006).
In ntrc plants, starch metabolism is impaired since particularly the accumulation of
starch was significantly lower when compared to wild type plants (Paper III).
However, starch remobilization was less affected in ntrc plants, suggesting that NTRC
preferably regulates starch synthesis than degradation in chloroplasts. NTRC has been
shown to monomerize the small subunits of AGPase in vitro in the presence of
NADPH (Michalska, et al., 2009). The yeast two-hybrid analysis (Paper III) confirmed
the interaction between AGPase and the thioredoxin domain of NTRC. On the
contrary, no interaction between NTRC domains and DSP4 was observed in yeast twohybrid assay. These results, together with the observation of the more severe effect of
NTRC knockout on starch synthesis rate than starch degradation rate in ntrc leaves,
attest the hypothesis that NTRC controls the starch synthesis but not the initial steps of
starch mobilization in Arabidopsis. However, NTRC may still be involved in
regulating other redox-controlled enzymes in starch degradation, since the maximal
apparent starch degradation rate was lower in ntrc plants when compared to wild type
plants (Paper III).
5.2.2. Altered homeostasis of metabolites synthesized from the end products of
shikimate acid pathway
The shikimate pathway, present only in plants and micro-organisms, links metabolism
of carbohydrates to biosynthesis of aromatic compounds. In a sequence of seven
metabolic steps, phosphoenolpyruvate and erythrose 4-phosphate are converted to
chorismate, the precursor of the aromatic amino acids tyrosin, tryptophan and
phenylalanine. These aromatic amino acids can then be used for protein synthesis or
they can be converted via several enzymatic steps to aromatic secondary metabolites
such as glucosinolates, phenylpropanoids including anthocyanins, and indole alkaloids
including growth hormone auxin (reviewed by Tzin and Galili, 2010).
The first enzyme in the pathway, 3-deoxy-d-arabino-heptulosonate-7-phosphate
synthase (DAHP synthase) which catalyzes the formation of 3-dehydroquianate from
phosphoenolpyruvate and erythrose 4-phosphate, has been reported to require reduced
thioredoxin for its activity (Entus, et al., 2002). Other enzymes in the pathway up to
chorismate are less well characterized and mechanisms that regulate their activity
remain to be elucidated. In ntrc plants the amounts of aromatic amino acids as well as
the amounts of auxin and anthocyanins were pronouncedly different from wild type
(Paper I). Shikimate-derived pathways are regulated by complex networks, including
feedback control by aromatic amino acids and environmental factors (Ishihara, et al.,
2007). Thus the lack of NTRC alters the homeostasis among the metabolic pathways
that derive from the shikimate pathway leading up to differential accumulation of
aromatic amino acids and to decreased accumulation of secondary metabolites auxin
and anthocyanins. Furthermore, in yeast two-hybrid analysis both DAHP synthase and
Discussion
47
tryptophan synthase β subunit gave specific interactions with thioredoxin domain of
NTRC (Figure 9), thus implicating that NTRC may be involved in regulating shikimate
pathway.
5.3. Thioredoxin- and ascorbate-dependent scavenging of ROS in
chloroplasts
5.3.1. Detoxification of H2O2 by NTRC
Thioredoxins participate in the avoidance of oxidative stress as they supply reducing
power to reductase enzymes involved in antioxidative metabolism, such as
peroxiredoxins (Rey, et al., 2005; Collin, et al., 2003; Finkemeier, et al., 2005) and
glutathione peroxidases (Herbette, et al., 2002; Jung, et al., 2002) (see 1.3.2.). The
reaction mechanism for peroxide reduction by peroxiredoxins involves a cysteine
residue that attacks the peroxide and becomes transiently oxidized to sulphenic acid.
This intermediate is subsequently attacked by the second cysteine residue yielding
water or the corresponding alcohol, and the two cysteine residues become oxidized and
form a disulphide bridge that needs to be reduced for a new catalytic cycle (Dietz,
2003; Konig, et al., 2003; Hall, et al., 2009). In chloroplasts, two pathways have been
proposed to reduce 2-Cys Prxs; one consists of FTR and free plastidial thioredoxins of
which Trx x is the most efficient (Collin, et al., 2003), another is based on NTRC
(Perez-Ruiz, et al., 2006; Kirchsteiger, et al., 2009; Perez-Ruiz and Cejudo, 2009).
Recently Pulido et al. (2010) demonstrated that NTRC is in fact the most relevant
pathway for chloroplast 2-Cys Prx reduction in vivo. The interaction between NTRC
and 2-Cys PrxB was confirmed also by the yeast two-hybrid analysis in Paper III.
Knockout of one antioxidative system in chloroplasts has demonstrated that the
detoxification of ROS never relies only on a single mechanism (Papers III and IV). The
decrease in the activity of NTRC/2-Cys Prxs was compensated by the elevation of
ascorbate-dependent antioxidative system in ntrc plants (Paper III; Pulido, et al., 2010).
Similarly, high-light-acclimated double mutant plants lacking both tAPX and sAPX
upregulated the accumulation of 2-Cys Prxs (Paper IV), indicating that chloroplast
APX-dependent and NTRC/2-Cys Prx-dependent systems are compensatory in the
detoxification of H2O2. Accordingly, plants deficient in NTRC do not accumulate
excess amount of ROS in leaf cells when compared to wild type (Paper III).
Nevertheless, the photoinhibition measurements, the activation state of malate
dehydrogenase and the carbonylation of proteins speak for the moderate oxidative
stress in ntrc leaves (Papers I and III; Pulido et al. 2010), indicating that ascorbatedependent detoxification system could not entirely balance the ROS metabolism in
chloroplasts. The total elimination of one antioxidant system may generally modify the
homeostasis of antioxidants in chloroplast and thereby impair the redox regulation of
metabolic enzymes.
48
Discussion
5.3.2. Impacts of chloroplast APXs on chloroplast redox homeostasis
Chloroplastic ascorbate peroxidases localized in the thylakoid membrane (tAPX) and
stroma (sAPX) have been unequivocally considered to play an essential role in
scavenging H2O2 in chloroplasts (Asada, 1999). The oxidation of the chloroplast APXs
by high concentration of H2O2 (Miyake, et al., 2006) has attenuated their physiological
significance regarding the defence response to photo-oxidative stress. In Paper IV, the
physiological function of tAPX and sAPX is clarified by characterization of single tapx
and sapx, and double tapx sapx lines. It is demonstrated that chloroplast APXs are
crucial in the protection against oxidative stress during greening of seedlings and upon
the short-term and long-term oxidative stress in mature leaves.
During seed germination, the mobilization of food storage by oxidative
phosphorylation in mitochondria generates ROS. In addition, light absorption of
chlorophylls (Hideg, et al., 2001) and deficient coupling between the two photosystems
support ROS formation in chloroplasts (Hutin, et al., 2003). To protect the developing
seedlings against ROS, activation of the antioxidant systems takes place early during
seedling development. Photo-oxidative stress during germination led to bleaching of
chloroplasts in seedlings lacking sAPX and especially in plants lacking both sAPX and
tAPX, whereas plants lacking only tAPX were equally resistant to photo-oxidative
stress during germination as wild type plants (Paper IV). Thus chloroplast APXs
contribute to antioxidant defence during the greening process, sAPX being primarily
important. Previously, both sAPX and tAPX have been shown to accumulate in pea
etioplasts in darkness and to decrease during de-etiolation and greening process
(Kanervo, et al., 2008), depicting their importance for photoprotection in developing
chloroplasts. In addition, the expression of genes encoding chloroplast APXs and Prxs
is upregulated during the first days after germination in Arabidopsis (Pena-Ahumada,
et al., 2006). Also glutaredoxin (Kanervo, et al., 2008), SOD and glutathione
transferase (Yang, et al., 2007) have been shown to be highly expressed in etiolated
plants and down-regulated upon greening of seedlings in light. Thereby the proper
induction of antioxidant systems in germinated seeds is crucial for the survival of a
seedling.
In mature leaves, tAPX and sAPX appear to be functionally redundant, and exhibit a
key role in the maintenance of chloroplast functionality upon sudden onset of oxidative
stress (Paper IV). In contrast, in the course of long-term acclimation to various stress
conditions, the chloroplast APXs can be compensated by other components of the
chloroplast antioxidative system (Paper IV). Therefore the stress-acclimation involves
the operations of multiple H2O2 detoxification systems to efficiently control the H2O2
levels in chloroplasts (see Figure 6). It was demonstrated in this thesis that in the
absence of both tAPX and sAPX, plants are able to scavenge H2O2 up to a certain
level, but if the production of H2O2 suddenly exceeds the threshold level, the lack of
tAPX and sAPX is not fully compensated by other components of the antioxidant
Discussion
49
network, suggesting that chloroplast APXs are indispensable under short-term changes
in the oxidative state of chloroplast, e.g. by strong sun flecks in nature. Accordingly,
high-light-acclimation of the tapx sapx double mutant plants enhanced the level of 2Cys Prxs (Paper IV), that can compensate the APXs in regard to H2O2 detoxification
(Dietz, et al., 2006). All high-light-acclimated plants, wild type, tapx, sapx and tapx
sapx double mutant plants accumulated also cytoplasmic APX (Paper IV). These
results suggest that both chloroplastic Prxs and cytoplasmic APX are more potent in
terms of H2O2 removal than chloroplast APXs under condition of elevated H2O2
production.
5.4. Photoperiodic development of photosynthetic tissues
Plant development is controlled by a number of external factors, of which light
quantity and quality and the length of the daily light period are the most essential ones.
As described in Papers I-II, the length of the daily light period induces developmental
alterations in Arabidopsis that are comparable to changes induced by light intensity.
Like plants grown under high light intensity (reviewed by Kim et al., 2005), long-daygrown plants possess thick leaves, vertically elongated palisade mesophyll cells, and
small grana stacks in chloroplasts when compared to plants grown under short days.
The length of the photoperiod also induces alterations in the physiological state of the
leaves. As described in Paper III, plants grown under short photoperiods show higher
accumulation of ROS after switching the light on subsequent to the dark period. Also
thylakoids isolated from short-day-grown plants produce more ROS than thylakoids
isolated from long-day-grown plants (Paper III). In respect of components of
antioxidative systems, the amounts of PrxIIE, 2-Cys Prxs and chloroplast GPX only
slightly increase in short-day-grown plants, being not sufficient to eliminate the higher
accumulation of ROS (Paper III). Accordingly, no significant differences could be
detected in the total content of ascorbate and glutathione between the short-day-grown
and long-day-grown plants (Queval, et al., 2007). The elevated production of ROS and
the low production of compensatory antioxidants in plants grown under short days
(Paper III; Becker, et al., 2006) may suggest that instead of oxidative stress, ROS have
a regulatory role in short-day-grown plants balancing metabolic reactions and the
regulation of development. Interestingly, Becker et al. (2006) showed that in plants
grown under long days, the expression of genes encoding several antioxidative
enzymes, including tAPX and sAPX, is strongly increased. This also indicates that
ROS-mediated signalling is specific for short photoperiods since under long
photoperiods ROS is aimed to be removed. All these studies collectively indicate that
an interaction between the day length and redox-mediated acclimation signals exists in
plants.
Diurnal metabolism of starch is one example of redox-regulated metabolic pathways
that balances the carbon fixation, carbon storage and the mobilization of stored carbon
according to the plant growth rate. As shown in Paper III, short photoperiods reduce
the growth of Arabidopsis. Both enzymes involved in starch synthesis, AGPase, and
50
Discussion
several enzymes involved in starch degradation, GWD, DSP4 and β-amylase1 (BAM1)
have been shown to be under redox control (Mikkelsen, et al., 2005; Sokolov, et al.,
2006; Sparla, et al., 2006). The higher accumulation of ROS in the leaves of short-daygrown plants soon after the onset of light (Paper III) suggests that chloroplasts
encounter higher oxidative pressure under short photoperiods than under long
photoperiods. Higher oxidation state in chloroplasts challenges the thioredoxin systems
and interferes with the redox-regulation of enzymes thus decreasing their optimal
activities.
5.4.1. Impact of NTRC knockout on the growth of Arabidopsis
The pleiotropic phenotype of ntrc plants suggests that there is no single reason for
growth reduction in plants deficient in NTRC. I have shown in my thesis that NTRC
contributes to important developmental, metabolic and defence processes in
chloroplasts, such as plastid division, biosynthesis of chlorophyll, starch and amino
acids and detoxification of H2O2. The significant reduction in chloroplast number and
the extent of irregularly developed chloroplasts observed in ntrc cells is probably the
most concrete factor that depresses the growth of ntrc plants. In addition, imbalances in
the important metabolic pathways in chloroplasts, starch metabolism and shikimate
pathway that leads to production of aromatic amino acids and secondary metabolites,
result in disorders in the functions of chloroplasts thus diminishing their optimal
performance. As confirmed by several reports, NTRC is a primary reductant for 2-Cys
Prxs (Paper III , (Perez-Ruiz, et al., 2006; Kirchsteiger, et al., 2009; Perez-Ruiz and
Cejudo, 2009; Pulido, et al., 2010). Thus plants lacking NTRC lack also one
component of the ROS scavenging mechanism in chloroplast resulting in increased
susceptibility to oxidative stress. However, it is noteworthy that the double mutant
plants deficient in both 2-Cys Prx A and B did not show any phenotype significantly
different from that of the wild type plants (Pulido, et al., 2010), indicating that the
reduced growth of ntrc line is mainly due to the turn down of the other NTRC target
processes. Since the highest growth reduction in ntrc plants was observed under short
photoperiods, it suggests that the demand for reductive capacity in the developmental
and metabolic processes, mediated by NTRC, is increased especially under short
photoperiods.
5.5. Chloroplast-to-nucleus retrograde signaling
The prevailing condition in ntrc cells - the existence of heterogeneous types of
chloroplasts in a single mesophyll cell - poses extraordinary challenges for the
retrograde signalling from chloroplast to nucleus that controls the nuclear gene
expression (reviewed by Nott et al., 2006 and Kleine et al., 2009). In plant cells, all
chloroplasts are autonomous in regard to biogenesis and function and they
communicate with the nucleus separately from each other (Yu, et al., 2007).
Discussion
51
Heterogenous chloroplast population in ntrc cells (Paper II) may send contradictory
signals to nucleus depending on their functional status, thereby confusing the nuclearcontrolled developmental processes. Similar irregularities in chloroplasts have been
reported in other mutant lines of chloroplast proteins (Knappe, et al., 2003; Hricova, et
al., 2006; Yu, et al., 2007). The deterioration of morphological development may be
caused by heterogeneous signals from chloroplasts that interfere with the development
of young leaves.
Light is the most eminent external factor that controls the plant development in nature.
Light is perceived by a set of wavelength-specific photoreceptors that initiate
signalling cascades which ultimately lead to adjustments in gene expression. Both
photoperiodic and photomorphogenetic development are regulated by the
phytochromes, which perceive red and far-red light, and the cryptochromes, which
respond to blue and UVA light (Jiao, et al., 2007). Transcript levels of genes coding for
cryptochrome 2 and the far-red light-impaired response regulator FRS3 are strongly
down-regulated in ntrc (Paper I). We also demonstrated that that the blue and far-red
light-mediated signaling is affected in ntrc plants (Paper I). Whether the downregulation of genes encoding blue light receptor and far-red-signaling component is
caused by the signals derived from the irregularly differentiated plastid in ntrc cell,
remains to be elucidated.
The transcript levels of four chloroplast-located proteases are elevated in ntrc (Paper I).
In the course of plant development, the quality and quantity of plastid proteins need to
be carefully controlled by chaperones and proteases. Proteases in plastids have two
types of function; firstly, they are involved in protein maturation and secondly, they
degrade damaged and unnecessary proteins to free amino acids (reviewed by
Sakamoto, 2006). Oxidation of proteins which takes place in a close vicinity to the
major cellular sites of ROS production induces damages to proteins and is a major
contributor to protein degradation. It was recently suggested that rather than ROS
themselves, peptides deriving from proteolytic degradation of oxidized proteins would
act as secondary ROS messengers thus contributing to retrograde signalling during
oxidative stress (Møller and Sweetlove, 2010).
52
Concluding remarks
6. Concluding remarks
I have demonstrated in this thesis that the acclimation of Arabidopsis to short
photoperiods/long nights increases the accumulation of ROS. This, in turn, increases
the demand for reductive systems to maintain the activities of redox-controlled proteins
involved in metabolic and developmental processes. Plants deficient in NTRC show
photoperiod-dependent phenotypes with more severe reduction in growth rate and
chlorophyll accumulation under short photoperiods than under long photoperiods.
These findings provide evidence that NTRC is particularly important regulator of
enzyme activities in plants grown under short photoperiods. In this thesis work, NTRC
was identified to be involved in controlling metabolic processes including starch
metabolism and shikimate pathway. Regulation by NTRC can be achieved directly by
mediating redox regulation of intramolecular or intermolecular disulfide bridges of
enzymes, or alternatively by protecting enzymes from oxidation via removal of H2O2 in
conjunction with 2-Cys Prxs. This thesis work also demonstrated that chloroplast
APXs, SOD and 2-Cys Prxs are tightly linked up to prevent the detrimental
accumulation of ROS in plants. Furthermore, it was shown that NTRC is needed for
regular biogenesis of chloroplasts in plant cells. However, the precise impact of NTRC
on the assembly and function of chloroplast division machinery and/or on the enzymes
of chlorophyll biosynthesis needs to be further clarified experimentally. Also the
search for novel NTRC target proteins by screening of the cDNA library with NTRC
baits is currently under process.
Acknowledgements
53
Acknowledgements
This work was carried out in the Laboratory of Molecular Plant Biology at the
University of Turku. Financial support from the Academy of Finland, the Finnish
Graduate School in Plant Biology and the Turku University Foundation is gratefully
acknowledged. My supervisor, Professor Eevi Rintamäki is warmly thanked for giving
me the opportunity to work in this project, and for excellent supervision and support in
scientific, as well as in non-scientific matters. Professor Eva-Mari Aro is thanked for
providing excellent working facilities in Biocity, and for her valuable scientific advice
and encouragement throughout these years.
Professors Elina Oksanen and Anja Hohtola are acknowledged for critically reviewing
this thesis.
I wish to acknowledge all my co-authors for excellent collaboration. Above all of the
previous and present co-workers I want to thank Saijaliisa Kangasjärvi for sharing her
knowledge and instructing me in the practical laboratory work with the peculiar
organism called Arabidopsis. Thank you Saija for letting me “watch you working”!
Ulla-Maija Suoranta and Mirja Jaala are acknowledged for keeping the laboratory
working in Biocity. I also want to thank Mika Keränen and Kurt Ståhle for their
invaluable help with all kinds of technical issues, and especially warm thanks are given
to Kurt who has patiently programmed the growth chambers to various light-dark
rhythms more than gazillion times.
“Transient group members” Ninni and Eve are warmly thanked for their support and
friendship. I am also grateful to everybody who has worked with me in Biocity during
these years; without you the working environment wouldn’t have been as pleasant and
fun and creative and supportive!
Of people outside the laboratory, enormous thanks are given to my sister Ullariikka
and her family for their love and encouragement. I also wish to thank my friends for
their support. And last, but definitely not least - Mika, thank you for being there for me
as we once promised to each other, “for better or for worse, and in sickness and in
health”.
Kiitos!
Kaarina July 2011
54
References
References
Akif M, Suhre K, Verma C, Mande SC (2005)
Conformational
flexibility
of
Mycobacterium
tuberculosis thioredoxin reductase: crystal structure
and normal-mode analysis. Acta Crystallogr D Biol
Crystallogr 61: 1603-1611
Balmer Y, Koller A, del Val G, Manieri W,
Schurmann P, Buchanan BB (2003) Proteomics
gives insight into the regulatory function of
chloroplast thioredoxins. Proc Natl Acad Sci U S A
100: 370-375
Allen JF, Puthiyaveetil S, Strom J, Allen CA (2005)
Energy transduction anchors genes in organelles.
Bioessays 27: 426-435
Balmer Y, Vensel WH, Hurkman WJ, Buchanan BB
(2006) Thioredoxin target proteins in chloroplast
thylakoid membranes. Antioxid Redox Signal 8:
1829-1834
Ankele E, Kindgren P, Pesquet E, Strand A (2007) In
vivo visualization of Mg-protoporphyrin IX, a
coordinator of photosynthetic gene expression in the
nucleus and the chloroplast. Plant Cell 19: 1964-1979
Apel K, Hirt H (2004) Reactive oxygen species:
metabolism, oxidative stress, and signal transduction.
Annu Rev Plant Biol 55: 373-399
Arsova B, Hoja U, Wimmelbacher M, Greiner E,
Ustun S, Melzer M, Petersen K, Lein W, Bornke F
(2010) Plastidial thioredoxin z interacts with two
fructokinase-like proteins in a thiol-dependent
manner: evidence for an essential role in chloroplast
development in Arabidopsis and Nicotiana
benthamiana. Plant Cell 22: 1498-1515
Asada K (1999) THE WATER-WATER CYCLE IN
CHLOROPLASTS: Scavenging of Active Oxygens
and Dissipation of Excess Photons. Annu Rev Plant
Physiol Plant Mol Biol 50: 601-639
Asada K, Kiso K, Yoshikawa K (1974) Univalent
reduction of molecular oxygen by spinach chloroplasts
on illumination. J Biol Chem 249: 2175-2181
Aseeva E, Ossenbuhl F, Sippel C, Cho WK, Stein B,
Eichacker LA, Meurer J, Wanner G, Westhoff P,
Soll J, Vothknecht UC (2007) Vipp1 is required for
basic thylakoid membrane formation but not for the
assembly of thylakoid protein complexes. Plant
Physiol Biochem 45: 119-128
Baena-Gonzalez E, Allahverdiyeva Y, Svab Z, Maliga
P, Josse EM, Kuntz M, Maenpaa P, Aro EM (2003)
Deletion of the tobacco plastid psbA gene triggers an
upregulation of the thylakoid-associated NAD(P)H
dehydrogenase complex and the plastid terminal
oxidase (PTOX). Plant J 35: 704-716
Baier M, Dietz KJ (2005) Chloroplasts as source and
target of cellular redox regulation: a discussion on
chloroplast redox signals in the context of plant
physiology. J Exp Bot 56: 1449-1462
Ballicora MA, Frueauf JB, Fu Y, Schurmann P,
Preiss J (2000) Activation of the potato tuber ADPglucose pyrophosphorylase by thioredoxin. J Biol
Chem 275: 1315-1320
Battchikova N, Vainonen JP, Vorontsova N, Keranen
M, Carmel D, Aro EM (2010) Dynamic changes in
the proteome of Synechocystis 6803 in response to
CO(2) limitation revealed by quantitative proteomics.
J Proteome Res 9: 5896-5912
Bauer J, Chen K, Hiltbunner A, Wehrli E, Eugster
M, Schnell D, Kessler F (2000) The major protein
import receptor of plastids is essential for chloroplast
biogenesis. Nature 403: 203-207
Bechtold U, Richard O, Zamboni A, Gapper C,
Geisler M, Pogson B, Karpinski S, Mullineaux PM
(2008) Impact of chloroplastic- and extracellularsourced ROS on high light-responsive gene expression
in Arabidopsis. J Exp Bot 59: 121-133
Beck CF (2005) Signaling pathways from
chloroplast to the nucleus. Planta 222: 743-756
the
Becker B, Holtgrefe S, Jung S, Wunrau C,
Kandlbinder A, Baier M, Dietz KJ, Backhausen
JE, Scheibe R (2006) Influence of the photoperiod on
redox regulation and stress responses in Arabidopsis
thaliana L. (Heynh.) plants under long- and short-day
conditions. Planta 224: 380-393
Benning C (2009) Mechanisms of lipid transport
involved in organelle biogenesis in plant cells. Annu
Rev Cell Dev Biol 25: 71-91
Blokhina O, Virolainen E, Fagerstedt KV (2003)
Antioxidants, oxidative damage and oxygen
deprivation stress: a review. Ann Bot 91 Spec No:
179-194
Blomqvist LA, Ryberg M, Sundqvist C (2008)
Proteomic analysis of highly purified prolamellar
bodies reveals their significance in chloroplast
development. Photosynth Res 96: 37-50
Boldt R, Zrenner R (2003) Purine and pyrimidine
biosynthesis in higher plants. Physiol Plant 117: 297304
Capitani G, Markovic-Housley Z, DelVal G, Morris
M, Jansonius JN, Schurmann P (2000) Crystal
structures of two functionally different thioredoxins in
spinach chloroplasts. J Mol Biol 302: 135-154
References
Cashmore AR, Jarillo JA, Wu YJ, Liu D (1999)
Cryptochromes: blue light receptors for plants and
animals. Science 284: 760-765
Chiadmi M, Navaza A, Miginiac-Maslow M, Jacquot
JP, Cherfils J (1999) Redox signalling in the
chloroplast: structure of oxidized pea fructose-1,6bisphosphate phosphatase. EMBO J 18: 6809-6815
Chibani K, Wingsle G, Jacquot JP, Gelhaye E,
Rouhier N (2009) Comparative genomic study of the
thioredoxin family in photosynthetic organisms with
emphasis on Populus trichocarpa. Mol Plant 2: 308322
Cho YH, Yoo SD, Sheen J (2006) Regulatory functions
of nuclear hexokinase1 complex in glucose signaling.
Cell 127: 579-589
Collin V, Issakidis-Bourguet E, Marchand C,
Hirasawa M, Lancelin JM, Knaff DB, MiginiacMaslow M (2003) The Arabidopsis plastidial
thioredoxins: new functions and new insights into
specificity. J Biol Chem 278: 23747-23752
Coudevylle N, Thureau A, Hemmerlin C, Gelhaye E,
Jacquot JP, Cung MT (2005) Solution structure of a
natural CPPC active site variant, the reduced form of
thioredoxin h1 from poplar. Biochemistry 44: 20012008
Dai S, Saarinen M, Ramaswamy S, Meyer Y, Jacquot
JP, Eklund H (1996) Crystal structure of Arabidopsis
thaliana NADPH dependent thioredoxin reductase at
2.5 A resolution. J Mol Biol 264: 1044-1057
Dat J, Vandenabeele S, Vranova E, Van Montagu M,
Inze D, Van Breusegem F (2000) Dual action of the
active oxygen species during plant stress responses.
Cell Mol Life Sci 57: 779-795
55
heptulosonate 7-phosphate synthase. Plant Physiol
129: 1866-1871
Fahnenstich H, Scarpeci TE, Valle EM, Flugge UI,
Maurino VG (2008) Generation of hydrogen
peroxide
in
chloroplasts
of
Arabidopsis
overexpressing glycolate oxidase as an inducible
system to study oxidative stress. Plant Physiol 148:
719-729
Falbel TG, Staehelin LA (1994) Characterization of a
family of chlorophyll-deficient wheat (Triticum) and
barley (Hordeum vulgare) mutants with defects in the
magnesium-insertion step of chlorophyll biosynthesis.
Plant Physiol 104: 639-648
Falini G, Fermani S, Ripamonti A, Sabatino P, Sparla
F, Pupillo P, Trost P (2003) Dual coenzyme
specificity of photosynthetic glyceraldehyde-3phosphate dehydrogenase interpreted by the crystal
structure of A4 isoform complexed with NAD.
Biochemistry 42: 4631-4639
Farquhar GD, von Caemmerer S, Berry JA (1980) A
biochemical model of photosynthetic CO2 assimilation
in leaves of C3 species. Planta 149, 1: 78-90
Fermani S, Ripamonti A, Sabatino P, Zanotti G,
Scagliarini S, Sparla F, Trost P, Pupillo P (2001)
Crystal structure of the non-regulatory A(4 )isoform of
spinach
chloroplast
glyceraldehyde-3-phosphate
dehydrogenase complexed with NADP. J Mol Biol
314: 527-542
Finkemeier I, Goodman M, Lamkemeyer P,
Kandlbinder A, Sweetlove LJ, Dietz KJ (2005) The
mitochondrial type II peroxiredoxin F is essential for
redox homeostasis and root growth of Arabidopsis
thaliana under stress. J Biol
Chem 280: 12168-12180
Desikan R, Cheung MK, Bright J, Henson D,
Hancock JT, Neill SJ (2004) ABA, hydrogen
peroxide and nitric oxide signalling in stomatal guard
cells. J Exp Bot 55: 205-212
Dietz KJ (2003) Plant peroxiredoxins. Annu Rev Plant
Biol 54: 93-107
Dietz KJ, Jacob S, Oelze ML, Laxa M, Tognetti V, de
Miranda SM, Baier M, Finkemeier I (2006) The
function of peroxiredoxins in plant organelle redox
metabolism. J Exp Bot 57: 1697-1709
Eggink LL, LoBrutto R, Brune DC, Brusslan J,
Yamasato A, Tanaka A, Hoober JK (2004)
Synthesis of chlorophyll b: localization of
chlorophyllide a oxygenase and discovery of a stable
radical in the catalytic subunit. BMC Plant Biol 4: 5
Entus R, Poling M, Herrmann KM (2002) Redox
regulation of Arabidopsis 3-deoxy-D-arabino-
Foyer CH, Noctor G (2009) Redox regulation in
photosynthetic organisms: signaling, acclimation, and
practical implications. Antioxid Redox Signal 11:
861-905
Foyer CH, Lopez-Delgado H, Dat JF, Scott IM (1997)
Hydrogen peroxide- and glutathione-associated
mechanisms of acclimatory stress tolerance and
signalling. Physiol Plantarum 100: 241-254
Galili G (2002) New insights into the regulation and
functional significance of lysine metabolism in plants.
Annu Rev Plant Biol 53: 27-43
Gao H, Sage TL, Osteryoung KW (2006) FZL, an
FZO-like protein in plants, is a determinant of
thylakoid and chloroplast morphology. Proc Natl Acad
Sci U S A 103: 6759-6764
Geck MK, Larimer FW, Hartman FC (1996)
Identification of residues of spinach thioredoxin f that
56
References
influence interactions with target enzymes. J Biol
Chem 271: 24736-24740
Gelhaye E, Rouhier N, Navrot N, Jacquot JP (2005)
The plant thioredoxin system. Cell Mol Life Sci 62:
24-35
Gibon Y, Blasing OE, Palacios-Rojas N, Pankovic D,
Hendriks JH, Fisahn J, Hohne M, Gunther M, Stitt
M (2004) Adjustment of diurnal starch turnover to
short days: depletion of sugar during the night leads to
a temporary inhibition of carbohydrate utilization,
accumulation of sugars and post-translational
activation of ADP-glucose pyrophosphorylase in the
following light period. Plant J 39: 847-862
Gill SS, Tuteja N (2010) Reactive oxygen species and
antioxidant machinery in abiotic stress tolerance in
crop plants. Plant Physiol Biochem 48: 909-930
Glynn JM, Miyagishima SY, Yoder DW, Osteryoung
KW, Vitha S (2007) Chloroplast division. Traffic 8:
451-461
Glynn JM, Froehlich JE, Osteryoung KW (2008)
Arabidopsis ARC6 coordinates the division
machineries of the inner and outer chloroplast
membranes through interaction with PDV2 in the
intermembrane space. Plant Cell 20: 2460-2470
Goslings D, Meskauskiene R, Kim C, Lee KP, Nater
M, Apel K (2004) Concurrent interactions of heme
and FLU with Glu tRNA reductase (HEMA1), the
target of metabolic feedback inhibition of tetrapyrrole
biosynthesis, in dark- and light-grown Arabidopsis
plants. Plant J 40: 957-967
Guo R, Luo M, Weinstein JD (1998) Magnesiumchelatase from developing pea leaves. Plant
Physiology 116: 605-615
Gutierrez-Nava MdlL, Gillmor CS, Jimenez LF,
Guevara-Garcia A, Leon P (2004) CHLOROPLAST
BIOGENESIS genes act cell and noncell
autonomously in early chloroplast development. Plant
Physiol 135: 471-482
Hägglund P, Bunkenborg J, Maeda K, Svensson B
(2008) Identification of thioredoxin disulfide targets
using a quantitative proteomics approach based on
isotope-coded affinity tags. J Proteome Res 7: 52705276
Hall A, Karplus PA, Poole LB (2009) Typical 2-Cys
peroxiredoxins--structures, mechanisms and functions.
FEBS J 276: 2469-2477
Hendriks JH, Kolbe A, Gibon Y, Stitt M,
Geigenberger
P
(2003)
ADP-glucose
pyrophosphorylase is activated by posttranslational
redox-modification in response to light and to sugars
in leaves of Arabidopsis and other plant species. Plant
Physiol 133: 838-849
Herbette S, Lenne C, Leblanc N, Julien JL, Drevet
JR, Roeckel-Drevet P (2002) Two GPX-like proteins
from Lycopersicon esculentum and Helianthus annuus
are antioxidant enzymes with phospholipid
hydroperoxide glutathione peroxidase and thioredoxin
peroxidase activities. Eur J Biochem 269: 2414-2420
Hernandez HH, Jaquez OA, Hamill MJ, Elliott SJ,
Drennan CL (2008) Thioredoxin reductase from
Thermoplasma acidophilum: a new twist on redox
regulation. Biochemistry 47: 9728-9737
Hideg E, Kalai T, Hideg K, Vass I (1998)
Photoinhibition of photosynthesis in vivo results in
singlet oxygen production detection via nitroxideinduced fluorescence quenching in broad bean leaves.
Biochemistry 37: 11405-11411
Hideg EE, Ogawa K, Kalai T, Hideg K (2001) Singlet
oxygen imaging in Arabidopsis thaliana leaves under
photoinhibition by excess photosynthetically active
radiation. Physiol Plant 112: 10-14
Hodges M, Miginiac-Maslow M, Decottignies P,
Jacquot JP, Stein M, Lepiniec L, Cretin C, Gadal P
(1994) Purification and characterization of pea
thioredoxin f expressed in Escherichia coli. Plant Mol
Biol 26: 225-234
Hricova A, Quesada V, Micol JL (2006) The
SCABRA3 nuclear gene encodes the plastid RpoTp
RNA polymerase, which is required for chloroplast
biogenesis and mesophyll cell proliferation in
Arabidopsis. Plant Physiol 141: 942-956
Husek P (1998) Chloroformates in gas chromatography
as general purpose derivatizing agents. J Chromatogr
B Biomed Sci Appl 717: 57-91
Hutin C, Nussaume L, Moise N, Moya I, Kloppstech
K, Havaux M (2003) Early light-induced proteins
protect Arabidopsis from photooxidative stress. Proc
Natl Acad Sci U S A 100: 4921-4926
Ido K, Ifuku K, Yamamoto Y, Ishihara S, Murakami
A, Takabe K, Miyake C, Sato F (2009) Knockdown
of the PsbP protein does not prevent assembly of the
dimeric PSII core complex but impairs accumulation
of photosystem II
supercomplexes in tobacco. Biochim Biophys Acta
1787: 873-881
Ikegami A, Yoshimura N, Motohashi K, Takahashi S,
Romano PG, Hisabori T, Takamiya K, Masuda T
(2007) The CHLI1 subunit of Arabidopsis thaliana
magnesium chelatase is a target protein of the
chloroplast thioredoxin. J Biol Chem 282: 1928219291
Inskeep WP, Bloom PR (1985) Extinction coefficients
of chlorophyll a and b in N,N-dimethylformamide and
80% acetone. Plant Physiol 77: 483-485
References
Ishihara A, Matsuda F, Miyagawa H, Wakasa K
(2007) Metabolomics for metabolically manipulated
plants: effects of tryptophan overproduction.
Metabolomics 3: 319-334
Jiao Y, Lau OS, Deng XW (2007) Light-regulated
transcriptional networks in higher plants. Nat Rev
Genet 8: 217-230
Johansson K, Ramaswamy S, Saarinen M, LemaireChamley M, Issakidis-Bourguet E, MiginiacMaslow M, Eklund H (1999) Structural basis for
light activation of a chloroplast enzyme: the structure
of sorghum NADP-malate dehydrogenase in its
oxidized form. Biochemistry 38: 4319-4326
Joyard J, Ferro M, Masselon C, Seigneurin-Berny D,
Salvi D, Garin J, Rolland N (2009) Chloroplast
proteomics and the compartmentation of plastidial
isoprenoid biosynthetic pathways. Mol Plant 2: 11541180
Jung BG, Lee KO, Lee SS, Chi YH, Jang HH, Kang
SS, Lee K, Lim D, Yoon SC, Yun DJ, Inoue Y, Cho
MJ, Lee SY (2002) A Chinese cabbage cDNA with
high sequence identity to phospholipid hydroperoxide
glutathione peroxidases encodes a novel isoform of
thioredoxin-dependent peroxidase. J Biol Chem 277:
12572-12578
Kanamaru K, Tanaka K (2004) Roles of chloroplast
RNA polymerase sigma factors in chloroplast
development and stress response in higher plants.
Biosci Biotechnol Biochem 68: 2215-2223
57
Kleine T, Voigt C, Leister D (2009) Plastid signalling
to the nucleus: messengers still lost in the mists?
Trends Genet 25: 185-192
Knappe S, Lottgert T, Schneider A, Voll L, Flugge
UI, Fischer K (2003) Characterization of two
functional
phosphoenolpyruvate/phosphate
translocator (PPT) genes in Arabidopsis--AtPPT1 may
be involved in the provision of signals for correct
mesophyll development. Plant J 36: 411-420
Kobayashi Y, Kanesaki Y, Tanaka A, Kuroiwa H,
Kuroiwa T, Tanaka K (2009) Tetrapyrrole signal as
a cell-cycle coordinator from organelle to nuclear
DNA replication in plant cells. Proc Natl Acad Sci U
S A 106: 803-807
Kolbe A, Oliver SN, Fernie AR, Stitt M, van Dongen
JT, Geigenberger P (2006) Combined transcript and
metabolite profiling of Arabidopsis leaves reveals
fundamental effects of the thiol-disulfide status on
plant metabolism. Plant Physiol 141: 412-422
Konig J, Lotte K, Plessow R, Brockhinke A, Baier M,
Dietz KJ (2003) Reaction mechanism of plant 2-Cys
peroxiredoxin. Role of the C terminus and the
quaternary structure. J Biol Chem 278: 24409-24420
Kötting O, Kossmann J, Zeeman SC, Lloyd JR (2010)
Regulation of starch metabolism: the age of
enlightenment? Curr Opin Plant Biol 13: 320-328
Krieger-Liszkay A (2005) Singlet oxygen production in
photosynthesis. J Exp Bot 56: 337-346
Kanervo E, Singh M, Suorsa M, Paakkarinen V, Aro
E, Battchikova N, Aro EM (2008) Expression of
protein complexes and individual proteins upon
transition of etioplasts to chloroplasts in pea (Pisum
sativum). Plant Cell Physiol 49: 396-410
Kroll D, Meierhoff K, Bechtold N, Kinoshita M,
Westphal S, Vothknecht UC, Soll J, Westhoff P
(2001) VIPP1, a nuclear gene of Arabidopsis thaliana
essential for thylakoid membrane formation. Proc Natl
Acad Sci U S A 98: 4238-4242
Karpinski S, Reynolds H, Karpinska B, Wingsle G,
Creissen G, Mullineaux P (1999) Systemic signaling
and acclimation in response to excess excitation
energy in Arabidopsis. Science 284: 654-657
Kuriyan J, Krishna TS, Wong L, Guenther B, Pahler
A, Williams CH,Jr, Model P (1991) Convergent
evolution of similar function in two structurally
divergent enzymes. Nature 352: 172-174
Kim GT, Yano S, Kozuka T, Tsukaya H (2005)
Photomorphogenesis of leaves: shade-avoidance and
differentiation of sun and shade leaves. Photochem
Photobiol Sci 4: 770-774
Laloi C, Rayapuram N, Chartier Y, Grienenberger
JM, Bonnard G, Meyer Y (2001) Identification and
characterization of a mitochondrial thioredoxin system
in plants. Proc Natl Acad Sci U S A 98: 14144-14149
Kirchsteiger K, Pulido P, Gonzalez M, Cejudo FJ
(2009) NADPH Thioredoxin reductase C controls the
redox status of chloroplast 2-Cys peroxiredoxins in
Arabidopsis thaliana. Mol Plant 2: 298-307
Larkin RM, Alonso JM, Ecker JR, Chory J (2003)
GUN4, a regulator of chlorophyll synthesis and
intracellular signaling. Science 299: 902-906
Kirkensgaard KG, Hagglund P, Finnie C, Svensson
B, Henriksen A (2009) Structure of Hordeum vulgare
NADPH-dependent
thioredoxin
reductase
2.
Unwinding the reaction mechanism. Acta Crystallogr
D Biol Crystallogr 65: 932-941
Larkindale J, Knight MR (2002) Protection against
heat stress-induced oxidative damage in Arabidopsis
involves calcium, abscisic acid, ethylene, and salicylic
acid. Plant Physiol 128: 682-695
Lee JW, Helmann JD (2006) The PerR transcription
factor senses H2O2 by metal-catalysed histidine
oxidation. Nature 440: 363-367
58
References
Lennon BW, Williams CH,Jr, Ludwig ML (2000)
Twists in catalysis: alternating conformations of
Escherichia coli thioredoxin reductase. Science 289:
1190-1194
Lennon BW, Williams CH,Jr, Ludwig ML (1999)
Crystal structure of reduced thioredoxin reductase
from Escherichia coli: structural flexibility in the
isoalloxazine ring of the flavin adenine dinucleotide
cofactor. Protein Sci 8: 2366-2379
Li Z, Wakao S, Fischer BB, Niyogi KK (2009) Sensing
and responding to excess light. Annu Rev Plant Biol
60: 239-260
S, Shimada H, Ohta H, Takamiya K (2003)
Functional analysis of isoforms of NADPH:
protochlorophyllide oxidoreductase (POR), PORB and
PORC, in Arabidopsis thaliana. Plant Cell Physiol 44:
963-974
Mateo A, Funck D, Muhlenbock P, Kular B,
Mullineaux PM, Karpinski S (2006) Controlled
levels of salicylic acid are required for optimal
photosynthesis and redox homeostasis. J Exp Bot 57:
1795-1807
Lunn JE (2007) Compartmentation in plant metabolism.
J Exp Bot 58: 35-47
Matsumoto F, Obayashi T, Sasaki-Sekimoto Y, Ohta
H, Takamiya K, Masuda T (2004) Gene expression
profiling of the tetrapyrrole metabolic pathway in
Arabidopsis with a mini-array system. Plant Physiol
135: 2379-2391
Maeda K, Finnie C, Svensson B (2004) Cy5 maleimide
labelling for sensitive detection of free thiols in native
protein extracts: identification of seed proteins
targeted by barley thioredoxin h isoforms. Biochem J
378: 497-507
Mazzella MA, Cerdan PD, Staneloni RJ, Casal JJ
(2001) Hierarchical coupling of phytochromes and
cryptochromes reconciles stability and light
modulation
of
Arabidopsis
development.
Development 128: 2291-2299
Maeda K, Hägglund P, Björnberg O, Winther JR,
Svensson B (2010) Kinetic and thermodynamic
properties of two barley thioredoxin h isozymes,
HvTrxh1 and HvTrxh2. FEBS Lett 584: 3376-3380
Mehler AH (1951) Studies on reactions of illuminated
chloroplasts. I. Mechanism of the reduction of oxygen
and other Hill reagents. Arch Biochem Biophys 33:
65-77
Maliga P (1998) Two plastid RNA polymerases of
higher plants: an evolving story. Trends Plant Sci 3: 46
Meskauskiene R, Nater M, Goslings D, Kessler F, op
den Camp R, Apel K (2001) FLU: a negative
regulator of chlorophyll biosynthesis in Arabidopsis
thaliana. Proc Natl Acad Sci U S A 98: 12826-12831
Manchado M, Michan C, Pueyo C (2000) Hydrogen
peroxide activates the SoxRS regulon in vivo. J
Bacteriol 182: 6842-6844
Maple J, Aldridge C, Moller SG (2005) Plastid
division is mediated by combinatorial assembly of
plastid division proteins. Plant J 43: 811-823
Maple J, Møller SG (2007) Interdependency of
formation and localisation of the Min complex
controls symmetric plastid division. J Cell Sci 120:
3446-3456
Marchand C, Le Marechal P, Meyer Y, MiginiacMaslow M, Issakidis-Bourguet E, Decottignies P
(2004) New targets of Arabidopsis thioredoxins
revealed by proteomic analysis. Proteomics 4: 26962706
Martin W (2003) Gene transfer from organelles to the
nucleus: frequent and in big chunks. Proc Natl Acad
Sci U S A 100: 8612-8614
Martin W, Herrmann RG (1998) Gene transfer from
organelles to the nucleus: how much, what happens,
and Why? Plant Physiol 118: 9-17
Masuda T, Fusada N, Oosawa N, Takamatsu K,
Yamamoto YY, Ohto M, Nakamura K, Goto K,
Shibata D, Shirano Y, Hayashi H, Kato T, Tabata
Meurer J, Plucken H, Kowallik KV, Westhoff P
(1998) A nuclear-encoded protein of prokaryotic
origin is essential for the stability of photosystem II in
Arabidopsis thaliana. EMBO J 17: 5286-5297
Meyer Y, Siala W, Bashandy T, Riondet C, Vignols F,
Reichheld JP (2008) Glutaredoxins and thioredoxins
in plants. Biochimica et Biophysica Acta (BBA) Molecular Cell Research 1783: 589-600
Michalska J, Zauber H, Buchanan BB, Cejudo FJ,
Geigenberger P (2009) NTRC links built-in
thioredoxin to light and sucrose in regulating starch
synthesis in chloroplasts and amyloplasts. Proc Natl
Acad Sci U S A 106: 9908-9913
Mikkelsen R, Mutenda KE, Mant A, Schurmann P,
Blennow A (2005) Alpha-glucan, water dikinase
(GWD): a plastidic enzyme with redox-regulated and
coordinated catalytic activity and binding affinity.
Proc Natl Acad Sci U S A 102: 1785-1790
Mittler R (2002) Oxidative stress, antioxidants and
stress tolerance. Trends Plant Sci 7: 405-410
Mittler R, Vanderauwera S, Gollery M, Van
Breusegem F (2004) Reactive oxygen gene network
of plants. Trends Plant Sci 9: 490-498
References
Miyagishima SY, Kabeya Y (2010) Chloroplast
division: squeezing the photosynthetic captive. Curr
Opin Microbiol 13: 738-746
Miyagishima S, Froehlich JE, Osteryoung KW (2006)
PDV1 and PDV2 mediate recruitment of the dynaminrelated protein ARC5 to the plastid division site. Plant
Cell 18: 2517-2530
Miyake C, Shinzaki Y, Nishioka M, Horiguchi S,
Tomizawa K (2006) Photoinactivation of ascorbate
peroxidase in isolated tobacco chloroplasts: Galdieria
partita APX maintains the electron flux through the
water-water cycle in transplastomic tobacco plants.
Plant Cell Physiol 47: 200-210
Mochizuki N, Tanaka R, Grimm B, Masuda T,
Moulin M, Smith AG, Tanaka A, Terry MJ (2010)
The cell biology of tetrapyrroles: a life and death
struggle. Trends Plant Sci 15: 488-498
Mochizuki N, Tanaka R, Tanaka A, Masuda T,
Nagatani A (2008) The steady-state level of Mgprotoporphyrin IX is not a determinant of plastid-tonucleus signaling in Arabidopsis. Proc Natl Acad Sci
U S A 105: 15184-15189
Mochizuki N, Brusslan JA, Larkin R, Nagatani A,
Chory J (2001) Arabidopsis genomes uncoupled 5
(GUN5) mutant reveals the involvement of Mgchelatase H subunit in plastid-to-nucleus signal
transduction. Proceedings of the National Academy of
Sciences 98: 2053-2058
Møller IM, Sweetlove LJ (2010) ROS signalling-specificity is required. Trends Plant Sci 15: 370-374
Montrichard F, Alkhalfioui F, Yano H, Vensel WH,
Hurkman WJ, Buchanan BB (2009) Thioredoxin
targets in plants: the first 30 years. J Proteomics 72:
452-474
Motohashi K, Kondoh A, Stumpp MT, Hisabori T
(2001) Comprehensive survey of proteins targeted by
chloroplast thioredoxin. Proc Natl Acad Sci U S A 98:
11224-11229
Moulin M, McCormac AC, Terry MJ, Smith AG
(2008) Tetrapyrrole profiling in Arabidopsis seedlings
reveals that retrograde plastid nuclear signaling is not
due to Mg-protoporphyrin IX accumulation. Proc Natl
Acad Sci U S A 105: 15178-15183
Mubarakshina MM, Ivanov BN, Naydov IA, Hillier
W, Badger MR, Krieger-Liszkay A (2010)
Production and diffusion of chloroplastic H2O2 and
its implication to signalling. J Exp Bot 61: 3577-3587
Mustardy L, Garab G (2003) Granum revisited. A
three-dimensional model--where things fall into place.
Trends Plant Sci 8: 117-122
59
Nakayama M, Masuda T, Bando T, Yamagata H,
Ohta H, Takamiya K (1998) Cloning and expression
of the soybean chlH gene encoding a subunit of Mgchelatase and localization of the Mg2+ concentrationdependent ChlH protein within the chloroplast. Plant
Cell Physiol 39: 275-284
Neff MM, Chory J (1998) Genetic interactions between
phytochrome A, phytochrome B, and cryptochrome 1
during Arabidopsis development. Plant Physiol 118:
27-35
Neuhaus HE, Emes MJ (2000) Nonphotosynthetic
metabolism in plastids. Annu Rev Plant Physiol Plant
Mol Biol 51: 111-140
Neuhaus HE, Stitt M (1990) Control analysis of
photosynthate partitioning: Impact of reduced activity
of ADP-glucose pyrophosphorylase or plastid
phosphoglucomutase on the fluxes to starch and
sucrose in Arabidopsis thaliana (L.) Heynh. Planta
182: 445-454
Nott A, Jung HS, Koussevitzky S, Chory J (2006)
Plastid-to-nucleus retrograde signaling. Annu Rev
Plant Biol 57: 739-759
Osteryoung KW, Stokes KD, Rutherford SM,
Percival AL, Lee WY (1998) Chloroplast division in
higher plants requires members of two functionally
divergent gene families with homology to bacterial
ftsZ. Plant Cell 10: 1991-2004
Pena-Ahumada A, Kahmann U, Dietz KJ, Baier M
(2006) Regulation of peroxiredoxin expression versus
expression of Halliwell-Asada-Cycle enzymes during
early seedling development of Arabidopsis thaliana.
Photosynth Res 89: 99-112
Perez-Ruiz JM, Cejudo FJ (2009) A proposed reaction
mechanism for rice NADPH thioredoxin reductase C,
an enzyme with protein disulfide reductase activity.
FEBS Lett 583: 1399-1402
Perez-Ruiz JM, Gonzalez M, Spinola MC, Sandalio
LM, Cejudo FJ (2009) The quaternary structure of
NADPH thioredoxin reductase C is redox-sensitive.
Mol Plant 2: 457-467
Perez-Ruiz JM, Spinola MC, Kirchsteiger K, Moreno
J, Sahrawy M, Cejudo FJ (2006) Rice NTRC is a
high-efficiency redox system for chloroplast
protection against oxidative damage. Plant Cell 18:
2356-2368
Peterson FC, Lytle BL, Sampath S, Vinarov D, Tyler
E, Shahan M, Markley JL, Volkman BF (2005)
Solution structure of thioredoxin h1 from Arabidopsis
thaliana. Protein Sci 14: 2195-2200
Piippo M, Allahverdiyeva Y, Paakkarinen V,
Suoranta U, Battchikova N, Aro E (2006)
Chloroplast-mediated regulation of nuclear genes in
60
References
Arabidopsis thaliana in the absence of light stress.
Physiol Genomics 25: 142-152
Pilon M, Ravet K, Tapken W (2011) The biogenesis
and physiological function of chloroplast superoxide
dismutases. Biochimica et Biophysica Acta (BBA) Bioenergetics In Press, Corrected Proof:
Porra RJ, Thompson WA, Kriedemann PE (1989)
Determination of accurate extinction coefficients and
simultaneous equations for assaying chlorophylls a
and b extracted with four different solvents:
verification of the concentration of chlorophyll
standards by atomic absorption spectroscopy.
Biochimica et Biophysica Acta (BBA) - Bioenergetics
975: 384-394
Pulido P, Spinola MC, Kirchsteiger K, Guinea M,
Pascual MB, Sahrawy M, Sandalio LM, Dietz KJ,
Gonzalez M, Cejudo FJ (2010) Functional analysis
of the pathways for 2-Cys peroxiredoxin reduction in
Arabidopsis thaliana chloroplasts. J Exp Bot 61: 40434054
Pyke KA (1999) Plastid division and development. Plant
Cell 11: 549-556
Pätsikkä E, Kairavuo M, Sersen F, Aro E, Tyystjärvi
E (2002) Excess copper predisposes photosystem II to
photoinhibition in vivo by outcompeting iron and
causing decrease in leaf chlorophyll. Plant Physiol
129: 1359-1367
Queval G, Issakidis-Bourguet E, Hoeberichts FA,
Vandorpe M, Gakiere B, Vanacker H, MiginiacMaslow M, Van Breusegem F, Noctor G (2007)
Conditional oxidative stress responses in the
Arabidopsis photorespiratory mutant cat2 demonstrate
that redox state is a key modulator of daylengthdependent gene expression, and define photoperiod as
a crucial factor in the regulation of H2O2-induced cell
death. Plant J 52: 640-657
Reed JW, Nagatani A, Elich TD, Fagan M, Chory J
(1994) Phytochrome A and phytochrome B have
overlapping but distinct functions in Arabidopsis
development. Plant Physiol 104: 1139-1149
Reichheld JP, Meyer E, Khafif M, Bonnard G, Meyer
Y (2005) AtNTRB is the major mitochondrial
thioredoxin reductase in Arabidopsis thaliana. FEBS
Lett 579: 337-342
Reinbothe C, Pollmann S, Reinbothe S (2010) Singlet
oxygen signaling links photosynthesis to translation
and plant growth. Trends Plant Sci 15: 499-506
Rey P, Cuine S, Eymery F, Garin J, Court M,
Jacquot JP, Rouhier N, Broin M (2005) Analysis of
the proteins targeted by CDSP32, a plastidic
thioredoxin participating in oxidative stress responses.
Plant J 41: 31-42
Rintamäki E, Martinsuo P, Pursiheimo S, Aro EM
(2000) Cooperative regulation of light-harvesting
complex II phosphorylation via the plastoquinol and
ferredoxin-thioredoxin system in chloroplasts. Proc
Natl Acad Sci U S A 97: 11644-11649
Ritte G, Lloyd JR, Eckermann N, Rottmann A,
Kossmann J, Steup M (2002) The starch-related R1
protein is an alpha -glucan, water dikinase. Proc Natl
Acad Sci U S A 99: 7166-7171
Rolland F, Baena-Gonzalez E, Sheen J (2006) Sugar
sensing and signaling in plants: conserved and novel
mechanisms. Annu Rev Plant Biol 57: 675-709
Rouhier N, Jacquot JP (2002) Plant peroxiredoxins:
alternative hydroperoxide scavenging enzymes.
Photosynth Res 74: 259-268
Ruggiero A, Masullo M, Ruocco MR, Grimaldi P,
Lanzotti MA, Arcari P, Zagari A, Vitagliano L
(2009) Structure and stability of a thioredoxin
reductase from Sulfolobus solfataricus: a thermostable
protein with two functions. Biochim Biophys Acta
1794: 554-562
Sakamoto W (2006) Protein degradation machineries in
plastids. Annu Rev Plant Biol 57: 599-621
Schmelz EA, Engelberth J, Alborn HT, O'Donnell P,
Sammons M, Toshima H, Tumlinson JH,III (2003)
Simultaneous analysis of phytohormones, phytotoxins,
and volatile organic compounds in plants. Proc Natl
Acad Sci U S A 100: 10552-10557
Schmitz AJ, Glynn JM, Olson BJSC, Stokes KD,
Osteryoung KW (2009) Arabidopsis FtsZ2-1 and
FtsZ2-2 are functionally redundant, but FtsZ-based
plastid division is not essential for chloroplast
partitioning or plant growth and development.
Molecular Plant 2: 1211-1222
Schoefs B, Franck F (2003) Protochlorophyllide
reduction: mechanisms and evolutions. Photochem
Photobiol 78: 543-557
Schürmann P, Buchanan BB (2008) The
ferredoxin/thioredoxin
system
of
oxygenic
photosynthesis. Antioxid Redox Signal 10: 1235-1274
Schürmann P, Jacquot JP (2000) Plant thioredoxin
systems revisited. Annu Rev Plant Physiol Plant Mol
Biol 51: 371-400
Schwarz O, Schurmann P, Strotmann H (1997)
Kinetics and thioredoxin specificity of thiol
modulation of the chloroplast H+-ATPase. J Biol
Chem 272: 16924-16927
Serrato AJ, Perez-Ruiz JM, Spinola MC, Cejudo FJ
(2004) A novel NADPH thioredoxin reductase,
localized in the chloroplast, which deficiency causes
References
hypersensitivity to abiotic stress in Arabidopsis
thaliana. J Biol Chem 279: 43821-43827
Slesak I, Libik M, Karpinska B, Karpinski S,
Miszalski Z (2007) The role of hydrogen peroxide in
regulation of plant metabolism and cellular signalling
in response to environmental stresses. Acta Biochim
Pol 54: 39-50
Sokolov LN, Dominguez-Solis JR, Allary AL,
Buchanan BB, Luan S (2006) A redox-regulated
chloroplast protein phosphatase binds to starch
diurnally and functions in its accumulation. Proc Natl
Acad Sci U S A 103: 9732-9737
Solymosi K, Schoefs B (2010) Etioplast and etiochloroplast formation under natural conditions: the
dark side of chlorophyll biosynthesis in angiosperms.
Photosynth Res 105: 143-166
Sparla F, Costa A, Lo Schiavo F, Pupillo P, Trost P
(2006) Redox regulation of a novel plastid-targeted
beta-amylase of Arabidopsis. Plant Physiol 141: 840850
Stenbaek A, Hansson A, Wulff RP, Hansson M, Dietz
KJ, Jensen PE (2008) NADPH-dependent
thioredoxin reductase and 2-Cys peroxiredoxins are
needed for the protection of Mg-protoporphyrin
monomethyl ester cyclase. FEBS Lett 582: 2773-2778
Stenbaek A, Jensen PE (2010) Redox regulation of
chlorophyll biosynthesis. Phytochemistry 71: 853-859
Stenbaek A (2010) Regulation of chlorophyll
biosynthesis – from light-harvesting to signalling
compounds. PhD thesis. University of Copenhagen,
Copenhagen
Strand A, Asami T, Alonso J, Ecker JR, Chory J
(2003) Chloroplast to nucleus communication
triggered by accumulation of Mg-protoporphyrinIX.
Nature 421: 79-83
Stumpp MT, Motohashi K, Hisabori T (1999)
Chloroplast thioredoxin mutants without active-site
cysteines facilitate the reduction of the regulatory
disulphide bridge on the gamma-subunit of chloroplast
ATP synthase. Biochem J 341 ( Pt 1): 157-163
Tanaka A, Tanaka R (2006) Chlorophyll metabolism.
Curr Opin Plant Biol 9: 248-255
Telfer A, Bishop SM, Phillips D, Barber J (1994)
Isolated photosynthetic reaction center of photosystem
II as a sensitizer for the formation of singlet oxygen.
Detection and quantum yield determination using a
chemical trapping technique. J Biol Chem 269:
13244-13253
The Arabidopsis Genome Initiative (2000) Analysis of
the genome sequence of the flowering plant
Arabidopsis thaliana. Nature 408: 796-815
61
Toledano MB, Delaunay A, Monceau L, Tacnet F
(2004) Microbial H2O2 sensors as archetypical redox
signaling modules. Trends Biochem Sci 29: 351-357
Tzin V, Galili G (2010) New insights into the shikimate
and aromatic amino acids biosynthesis pathways in
plants. Mol Plant 3: 956-972
Ursini F, Maiorino M, Brigelius-Flohe R, Aumann
KD, Roveri A, Schomburg D, Flohe L (1995)
Diversity of glutathione peroxidases. Methods
Enzymol 252: 38-53
Veljovic-Jovanovic S, Noctor G, Foyer CH (2002) Are
leaf hydrogen peroxide concentrations commonly
overestimated? The potential influence of artefactual
interference by tissue phenolics and ascorbate. Plant
Physiology and Biochemistry 40: 501-507
Verdoucq L, Vignols F, Jacquot JP, Chartier Y,
Meyer Y (1999) In vivo characterization of a
thioredoxin h target protein defines a new
peroxiredoxin family. J Biol Chem 274: 19714-19722
Vignols F, Brehelin C, Surdin-Kerjan Y, Thomas D,
Meyer Y (2005) A yeast two-hybrid knockout strain
to explore thioredoxin-interacting proteins in vivo.
Proc Natl Acad Sci U S A 102: 16729-16734
Vothknecht UC, Westhoff P (2001) Biogenesis and
origin of thylakoid membranes. Biochimica et
Biophysica Acta (BBA) - Molecular Cell Research
1541: 91-101
Waksman G, Krishna TS, Williams CH,Jr, Kuriyan J
(1994) Crystal structure of Escherichia coli
thioredoxin reductase refined at 2 A resolution.
Implications for a large conformational change during
catalysis. J Mol Biol 236: 800-816
Wang Q, Sullivan RW, Kight A, Henry RL, Huang J,
Jones AM, Korth KL (2004) Deletion of the
chloroplast-localized Thylakoid Formation1 gene
product in Arabidopsis leads to deficient thylakoid
formation and variegated leaves. Plant Physiol 136:
3594-3604
Weber A, Flugge UI (2002) Interaction of cytosolic and
plastidic nitrogen metabolism in plants. J Exp Bot 53:
865-874
Willekens H, Chamnongpol S, Davey M, Schraudner
M, Langebartels C, Van Montagu M, Inze D, Van
Camp W (1997) Catalase is a sink for H2O2 and is
indispensable for stress defence in C3 plants. EMBO J
16: 4806-4816
Wolosiuk RA, Buchanan BB (1978) Activation of
chloroplast NADP-linked glyceraldehyde-3-phosphate
dehydrogenase by the ferredoxin/thioredoxin system.
Plant Physiology 61: 669-671
62
References
Woodson JD, Chory J (2008) Coordination of gene
expression between organellar and nuclear genomes.
Nat Rev Genet 9: 383-395
Yamasato A, Nagata N, Tanaka R, Tanaka A (2005)
The N-terminal domain of chlorophyllide a oxygenase
confers protein instability in response to chlorophyll B
accumulation in Arabidopsis. Plant Cell 17: 15851597
Yang P, Chen H, Liang Y, Shen S (2007) Proteomic
analysis of de-etiolated rice seedlings upon exposure
to light. Proteomics 7: 2459-2468
Yang Y, Glynn JM, Olson BJ, Schmitz AJ,
Osteryoung KW (2008) Plastid division: across time
and space. Curr Opin Plant Biol 11: 577-584
Yano S, Terashima I (2001) Separate localization of
light signal perception for sun or shade type
chloroplast and palisade tissue differentiation in
Chenopodium album. Plant Cell Physiol 42: 13031310
Yu F, Fu A, Aluru M, Park S, Xu Y, Liu H, Liu X,
Foudree A, Nambogga M, Rodermel S (2007)
Variegation mutants and mechanisms of chloroplast
biogenesis. Plant Cell Environ 30: 350-365
Yu TS, Kofler H, Hausler RE, Hille D, Flugge UI,
Zeeman SC, Smith AM, Kossmann J, Lloyd J,
Ritte G, Steup M, Lue WL, Chen J, Weber A
(2001) The Arabidopsis sex1 mutant is defective in
the R1 protein, a general regulator of starch
degradation in plants, and not in the chloroplast
hexose transporter. Plant Cell 13: 1907-1918
Zeeman SC, Kossmann J, Smith AM (2010) Starch: its
metabolism,
evolution,
and
biotechnological
modification in plants. Annu Rev Plant Biol 61: 209234
Zeeman SC, Northrop F, Smith AM, Rees T (1998) A
starch-accumulating mutant of Arabidopsis thaliana
deficient in a chloroplastic starch-hydrolysing
enzyme. Plant J 15: 357-365
Zeeman SC, Smith SM, Smith AM (2007) The diurnal
metabolism of leaf starch. Biochem J 401: 13-28
Zhang N, Portis AR,Jr (1999) Mechanism of light
regulation of Rubisco: a specific role for the larger
Rubisco activase isoform involving reductive
activation by thioredoxin-f. Proc Natl Acad Sci U S A
96: 9438-9443
Zheng M, Aslund F, Storz G (1998) Activation of the
OxyR transcription factor by reversible disulfide bond
formation. Science 279: 1718-1721